Self-assembled metal colloid monolayers having size and density gradients

ABSTRACT

A biosensor based on complexes between biomolecule receptors and colloidal Au nanoparticles, and more specifically, colloid layers of receptor/Au complexes that can be used to detect biomolecule analytes through measuring of binding-induced changes in electrical resistance or surface plasmon resonance. Also disclosed is a method for detecting and analysing carrier-borne chemical compounds with Raman spectroscopy using an improved SERS substrate. Further disclosed is an improved method for detecting compounds in solvents using capillary electrophoresis in conjunction with Raman spectroscopy.

This is the National Stage of International Application No.PCT/US97/15581, filed Sep. 4, 1997; which is a continuation-in-part ofapplication Ser. No. 08/769,970, filed Dec. 19, 1996, now abandoned andwhich claims the benefit of U.S. Provisional Application No. 60/025,064,filed Sep. 4, 1996.

BACKGROUND OF THE INVENTION

The present invention relates to self-assembled metal colloidmonolayers, methods of preparation, and use thereof.

In surface enhanced Raman scattering (SERS), million-fold enhancementsin Raman scattering can be obtained for molecules adsorbed at suitablyrough surfaces of Au, Ag, and Cu. Although many approaches have beenreported, preparation of well-defined, stable SERS substrates havinguniform roughness on the critical 3 to 100 nm scale has provendifficult. Because colloidal Au can be synthesized as monodispersesolutions throughout most of this size regime, and because moleculesadsorbed to closely spaced colloidal Au and Ag exhibit enhanced Ramanscattering, these particles are excellent building blocks forSERS-active substrates. The key issue is whether colloidal Au and Agparticles can be organized into macroscopic surfaces that have awell-defined and uniform nanometer-scale architecture. Indeed,controlling nanostructure is currently a central focus throughoutmaterials research. Progress in self assembly of organic thin films onmetal surfaces [C. D. Bain and G. M. Whitesides, Angew. Chem. Int. EdEngl. 28, 506 (1989); A. Ulman, An Introduction to Ultrathin OrganicFilms, from Langmuir-Blodgett to Self-Assembly (Academic Press, Boston,1991)] led us to explore the reverse process: self assembly of colloidalAu and Ag particles onto supported organic films. As detailed below,this approach has yielded surfaces that are SERS-active, characterizableat both the macroscopic and microscopic levels, highly reproducible,electrochemically addressable, and simple to prepare in large numbers.Moreover, these substrates have a surface roughness that is defined bythe colloid diameter (which is tunable) and an average interparticlespacing that is continuously variable. As such, self-assembled Au, Agand Ag-coated colloid monolayers are likely to have extraordinaryutility for SERS.

In the nearly twenty years since the discovery of surface enhanced Ramanscattering (SERS) of molecules adsorbed at roughened Ag electrodes, andthe accompanying theoretical work demonstrating the need for surfaceroughness, there have been numerous reports of new architectures forSERS substrates. See, for instance, Liao, P. F.; Bergman, J. G.; Chemla,D. S.; Wokaun, A.; Melngailis, J.; Hawyrluk, A. M.; Economou, N. P.Chem. Phys. Lett. 1981, 82, 355-9; Creighton, J. A.; Blatchford, C. G.;Albrecht, M. G. J. Chem. Soc., Faraday Trans. 2 1979, 75, 790-8;Blatchford, C. G.; Campbell, J. R.; Creighton, J. A. Surf Sci. 1982,120, 435-55; Tran, C. D. Anal. Chem. 1984, 56, 824-6; Soper, S. A.;Ratzhlaff, K. L.; Kuwana, T. Anal. Chem. 1990, 62, 143844; Sequaris,J.-M.; Koglin, E. Fresenius J Anal. Chem. 1985, 321, 758-9; Aroca, R.;Jennings, C.; Kovacs, G. J.; Loutfy, R. O.; Vincett, P. S. J. Phys.Chem. 1985, 89, 4051-4; Moody, R. L.; Vo-Dinh, T.; Fletcher, W. H. Appl.Spectrosc. 1987, 41, 966-70; Ni, F.; Cotton, T. M. Anal. Chem. 1986, 58,3159-63; Yogev, D.; Efrima, S. J. Phys. Chem. 1988, 92, 5761-5;Goudonnet, J. P.; Bijeon, J. L.; Warmack, R. J.; Ferrell, T. L. Phys.Rev. B: Condensed Matter 1991, 43, 4605-12; Murray, C. A.; Allara, D. L.J. Chem. Phys. 1982, 76, 1290-1303; Brandt, E. S. Appl. Spectrosc. 1993,47, 85-93; Alsmeyer, Y. W.; McCreery, R. L. Anal. Chem. 1991, 63,1289-95; Mullen, K.; Carron, K. Anal. Chem. 1994, 66, 478-83; Beer, K.D.; Tanner, W.; Garrell, R. L. J. Electroanal Chem. 1989, 258, 313-25;Dawson, P.; Alexander, K. B.; Thompson, J. R.; Haas III, J. W.; Ferrell,T. L. Phys. Rev. B: Condens. Matter 1991, 44, 6372-81; Roark, S. E.;Rowlen, K. L. Appl. Spectrosc. 1992, 46, 1759-61; Roark, Shane E.;Rowlen, K. L. Chem. Phys. Lett. 1993, 212, 50; Roark, Shane E.; Rowlen,K. L. Anal. Chem. 1994, 66, 261-70; Walls, D.; Bohn, P. J. Phys. Chem.1989, 93, 2976-82; Dutta, P. K.; Robins, D. Langmuir 1991, 7, 2004-6;Sheng, R.-S.; Zhu, L.; Morris, M. D. Anal. Chem. 1986, 58,1116-9.

These surfaces span a wide range of assembly principles and encompasssimilarly broad levels of complexity. Examples of SERS-active surfacesinclude electrochemically-roughened electrodes,microlithographically-prepared elliptical Ag posts, aggregates ofcolloidal Au or Ag particles—both in solution and associated withchromatographic media, evaporated thin films, Ag-coated latex particles,substrates prepared by chemical reduction of Ag⁺, and liquid Ag films.The motivation for this work stems from several intrinsically attractiveaspects of SERS as a vibrational spectroscopy-based structural tooland/or analytical method: million fold signal enhancements compared tosolution Raman spectra, adsorption-induced fluorescence quenching, alack of interference from H₂O, and molecular generality. However, whileSERS has been invaluable for certain narrowly defined applications, mostspectroscopists would agree that the technique has not lived up to itsenormous potential.

The problem has been the inability of any previous surface to meet all,or even most, of the essential criteria that would define a truly usefulSERS substrate: strongly enhancing, reproducible, uniformly rough, easyto fabricate, and stable over time. Biocompatibility is also extremelyimportant, insofar as previous studies demonstrating partial or fullprotein denaturation upon adsorption to SERS-active substrates [Holt, R.E.; Cotton, T. M. J: Am. Chem. Soc. 1989, 111, 2815-21; Lee, N.-S.;Hsieh, Y.-Z.; Morris, M. D.; Schopfer, L. M. J. Am. Chem. Soc. 1987,109, 1353-63] have proven to be a major setback to the use of SERS inbiological systems. Other desirable characteristics includeelectromagnetic tunability (i.e. the ability to control the wavelengthwhere optimal enhancement occurs, so as to match the substrate to thephoton source), electrochemical addressability—to control the extent ofadsorption and the redox state of adsorbed species, a lack of surface“activation” steps, and a low cost per substrate. This last feature isparticularly important for applications involving large numbers ofroutine measurements, such as in environmental monitoring, airportsecurity, and clinical medicine.

SUMMARY OF THE INVENTION

Detailed embodiments of the present invention are disclosed herein.However, it is understood that certain preferred embodiments are merelyillustrative of the invention which may be embodied in various forms andapplications. Specific compositional and functional details disclosedherein are not meant to be interpreted as limiting, but merely assupport for the invention as claimed and as appropriate representationsfor teaching those skilled in the art to variously employ the presentinvention in any appropriate embodiment.

We report here a new approach to SERS substrates that meets all of thecriteria delineated above. Our strategy involves assembly of colloidalAu, Ag, or other suitable metal particles into macroscopictwo-dimensional arrays on polymer-immobilized substrates (FIG. 1). Inthe first, covalent approach, reactive hydroxyl/oxide groups aregenerated on a substrate. For many substrates (glass, metal, etc.), suchfunctional groups are already present in high concentration. A secondstep involves surface-initiated polymerization of bifunctionalorganosilanes such as (RO)₃Si(CH₂)₃A. The alkoxysilane forms covalentattachments to the surface via hydrolysis. The pendant functional groupA (FIG. 1B), chosen for its high affinity toward noble metal surfaces,extends out into solution. In the final step, the polymer-derivatizedsubstrate is immersed into a solution of colloidal Au particles, wheresurface assembly spontaneously occurs. An alternate approach based onhigh affinity binding of streptavidin to biotin can also be used (FIG.1C). See Wilchek, M.; Bayer, E. A. Anal. Biochem. 1988, 171, 1 andAnzai, J.; Hoshi, T.; Osa, T Trends Anal. Chem. 1994, 13, 205-10. Here,a biotinylated surface is reacted with a colloidal Au-streptavidinconjugate to form a colloid-based surface held together by non-covalentinteractions.

With molecular self-assembly on metal substrates [Nuzzo, R. G.; Allara,D. L. J. Am. Chem. Soc. 1983, 105, 4481-4483; Bain, C. D.; Whitesides,G. M. Angew. Chem. Int. Ed. Engl. 1989, 28, 506-512] now established asan important route to controlling interfacial properties, it should bepointed out that the approaches delineated in FIG. 1 define what isessentially the inverse process: self-assembly of well-definedparticulate metal films on organic substrates. The term self-assemblyrefers to our finding that interparticle spacing is governed byinterparticle repulsive forces. We have explored this chemistry indetail by varying substrate, polymer, colloid diameter, and reactionconditions. Moreover, the electrochemical characteristics ofcolloid-based surfaces, the kinetics of surface formation, and theelectromagnetic properties of composite particles have beeninvestigated. We also describe here the basic steps involved in surfaceassembly and characterization, as well as experimental verification ofSERS activity. What distinguishes this work are the following features:macroscopic surfaces of controlled and uniform roughness can be preparedby self-assembly, the resulting substrates are compatible withbiomolecules, and the surfaces exhibit a high degree ofdurability/stability over time.

Our construction protocol for SERS-active Au and Ag colloid monolayersexploits the simplicity of self assembly from solution and the affinityof noble metal surfaces for certain organic functional groups (FIG. 1).In our case, these moieties are present by virtue of organic filmseither polymerized or deposited on the surface of macroscopic (0.8 cm×2cm) substrates. Immersion of the functionalized substrate into a dilutesolution of monodisperse colloidal Au or Ag particles leads to colloidimmobilization. This solution-based process is extremely general,encompassing numerous permutations of insulating and conductingsubstrates [glass, quartz, plasma-treated Teflon, Formvar, indium-dopedSnO₂ (ITO), and Pt], organic films [hydrolyzed mono-, di- andtrialkoxysilanes containing the functional groups CN, NH₂, 2-pyridyl,P(C₆H₅)₂, and SH, as well as carboxyl-terminated C₁₈ organothiolself-assembled monolayers], and colloids [5 to 70 nm in diameter for Au,and 5 to 20 nm in diameter for Ag and Au/Ag composites]. Our work hasfocused on Au and Ag particles, but with the right functional group A, awide variety of colloidal particles could constitute building blocks forwell-defined macroscopic surfaces.

Solution-based surface assembly also eliminates preparative, geometric,and operational constraints associated with most previously describedSERS substrates. Thus, one liter of 17 nM, 12-nm diameter colloidal Au,which can be stored indefinitely at room temperature, can be used toprepare 2,000 0.5-cm² surfaces with only a 1% decrease in colloidconcentration. Importantly, these substrates can be assembledsequentially or simultaneously. Surfaces in novel geometries that extendthe utility of SERS can now be derivatized, including one face of a 5ml-volume spectroelectrochemical cell, large glass sheets severalcentimeters on a side, and the inside of a 20-mm inner diameter glasscapillary. Moreover, once constructed, no further activation steps (suchas electrochemical oxidation-reduction cycles or particle aggregation)are required to initiate SERS activity. It should be noted that theparticles are tightly bound and the thermodynamic stability of thesesurfaces is very high: exchange with molecules in solution containingthe functional group A does not occur.

This preparation method differs greatly from electrochemical rougheningof electrodes and metal vapor deposition, the most common routes tosolid SERS substrates. Each of these protocols yields surfaces withpolydisperse roughness on the nanometer scale. This problem iscircumvented by the methods of FIG. 1. Since the size of the colloidprecursor can be easily varied and controlled, the defining roughness ofthe surface is pre-determined. Not only can the roughness be tunedaccording to experimental needs, but the roughness is uniform—allparticles are of the same size and dimensions. This is of particularimportance in SERS where enhancement at the surface is directlycorrelated to nanometer scale roughness.

Two-dimensional colloid self assembly also differs from the numerousmethods for preparation of SERS-active substrates involving colloidalparticles [Creighton, J. A.; Blatchford, C. G.; Albrecht, M. G. J. Chem.Soc., Faraday Trans. 2 1979, 75, 790-8; Blatchford, C. G.; Campbell, J.R.; Creighton, J. A. Surf Sci. 1982, 120, 435-55; Tran, C. D. Anal.Chem. 1984, 56, 824-6; Soper, S. A.; Ratzhlaff, K. L.; Kuwana, T. Anal.Chem. 1990, 62, 1438-44; Sequaris, J.-M.; Koglin, E. Fresenius J. Anal.Chem. 1985, 321, 758-9; Ahern, A. M.; Garrell, R. L. Langmuir 1991, 7,254-61; Angel, S. M.; Katz, L. F.; Archibald, D. D.; Honigs, D. E. Appl.Spectrosc. 1989, 43, 367-72; Clarkson, J.; Campbell, C.; Rospendowski,B. N.; Smith, W. E. J. Raman Spectrosc. 1991, 22, 771-775]. In thosemethods, there is a single size of particle, but since there is nocontrol over interparticle interactions, aggregates of ill-defineddimensions are often formed. In this work, strong covalent ornon-covalent bonds to the substrate reduce the surface mobility of thenanoparticles and prevent the spontaneous coalescence of particles onthe surface. Thus, the initial size uniformity is maintained.

For several reasons, keeping the particles physically separated is acritical component to our assembly strategy. (1) The intrinsicbiocompatibility of individual colloidal Au particles is maximized:aggregates begin to approximate larger surfaces where, for Au, proteindenaturation is a serious concern. (2) The resulting surfaces are morestraightforwardly characterized than particle aggregates. (3) It isknown both from theory and experiment that closely-spaced but physicallyseparated particle arrays can be strongly enhancing [Inoue, M.; Ohtaka,K. J. Phys. Soc. Jpn. 1983, 52, 3853-64; Chu, L.-C.; Wang, S.-Y. J.Appl. Phys. 1985, 57, 453-9; Chu, L.-C.; Wang, S.-Y. Phys. Rev. B:Condens. Matter 1985, 31, 693-9]. As the interparticle spacing increasestoward λ, nearly all of the SERS effect is lost: completely isolatedsmall colloidal Au particles are weakly enhancing. In our view, therelatively small loss in enhancement for non-contacting, closely-spacedparticles is more than offset by an increased ease of characterization,improved biocompatibility, and demonstrated improvements in stability(vide infra). FIG. 1D depicts various regimes for colloid immobilizationthat could result from using the strategy delineated above. In surfaceA, the particles are isolated, but too far apart to be stronglyenhancing. In B, the particles are close enough to see the SERS effect,but still isolated (thus retaining the biocompatibility properties ofindividual particles). Surface C represents a close-packed colloidmonolayer, while D represents particle multilayers approximating a bulksurface. The latter surface can be prepared from a monolayer. Our goalsare to prepare and characterize surfaces like B, C and D, and to usethem to solve a variety of problems in analytical, biological,environmental, clinical and inorganic chemistry.

To this end, the ease of fabrication and handling of Au colloidmonolayers is very significant. Large numbers of samples can be preparedsimultaneously, with no restrictions on the size or shape of thesubstrates, and without the need for even moderately sophisticatedequipment (i.e. no potentiostat, no vacuum deposition apparatus).Furthermore, with transparent substrates, the optical properties of theSERS-active surface can be monitored directly. This means that areasonably accurate prediction of enhancement factors can be made apriori. Indeed, as described below, uv-vis is our basic characterizationtool.

BRIEF DESCRIPTION OF THE FIGURES

FIGS. 1A-1B show assembly strategies for Au and Ag colloid monolayers;X═A═CN, NH₂, 2-pyridyl, P(C₆H₅)₂, and SH.

FIG. 1C demonstrates one embodiment of the present invention based onhigh affinity binding of streptavidin to a biotin.

FIG. 1D depicts various degrees of substrate surface coverage by metalparticles.

FIG. 2A illustrates the ultraviolet-visible kinetics of a Au colloidmonolayer on a glass substrate coated with3-aminopropyltrimethoxysilane.

FIG. 2B illustrates the SERS kinetics of a Au colloid monolayer on aglass substrate coated with 3-aminopropyltrimethoxysilane.

FIG. 3A is a TEM image of a Au colloid monolayer prepared on aSiO_(x)-coated formvar surface.

FIG. 3B shows the SERS spectrum of BPE adsorbed onto the derivatized TEMgrid used to produce FIG. 3A.

FIG. 4 shows an electrochemical potential dependence of SERS intensityof the 1006 cm⁻¹ band of pyridine on a Ag colloid monolayer on Pt and onbulk Ag.

FIG. 5 shows the optical and SERS spectra before and after deposition ofAg onto 18 nm diameter colloidal Au monolayer.

FIG. 6A shows optical spectra for solutions of isolated and aggregated13 nm colloidal Au particles in H₂O.

FIG. 6B illustrates the diversity of optical properties attainablethrough the method of the present invention.

FIG. 7 illustrates the optical stability of the Au colloid-basedmonolayers of the present invention.

FIG. 8 shows the optical spectrum for a MP-biocytin coated quartz slidecontaining a monolayer of Au derivatized with BSA and streptavidin.

FIGS. 9A-9B show TEM micrographs of a colloidal Au surface prepared bythe method of the present invention.

FIG. 9C shows a TEM micrograph of a formvar-coated TEM grid which wasfloated in colloidal Au for 2 days (not the method of the presentinvention).

FIG. 10A shows a SERS spectrum of BPE drop-coated on a colloid Aumonolayer on a functionalized TEM grid, as well as the SERS spectrum ofthe adsorbate-free surface.

FIG. 10B shows a SERS spectrum of BPE drop-coated on a colloid Aumonolayer on a HO₂C(CH₂)₁₆SH/Au/Cr/Si substrate, as well as the SERSspectrum of the adsorbate-free surface.

FIGS. 11-13 are TEM images of the colloidal particles prepared inaccordance with example 1.

FIG. 14 is the solution optical spectrum of the seed nuclei of example1.

FIGS. 15-16 are uv-vis spectra of the larger particles derivatized onglass substrates and of the larger particles in solution as described inexample 1.

FIG. 17 shows a TEM image of the colloidal surface in example 2 afterthe first layer of colloidal particles has been applied.

FIG. 18 shows an optical spectrum of the colloid surface in example 2after the first layer of colloidal particles has been applied.

FIG. 19 shows an optical spectrum of the colloid surface in example 2after the second layer of colloidal particles has been applied.

FIG. 20 shows non-covalent colloidal approaches to protein coated metalsurfaces.

FIG. 21 is a comparison of the optical properties of covalent andnon-covalent attachment.

FIG. 22 shows the changes in optical spectra upon electrochemicalreduction of Ag⁺ ions onto Au colloid monolayers in accordance withexample 4.

FIG. 23 shows theoretical modeling of the expected change in opticalspectra of Ag-clad Au colloid monolayers for increasing deposits of Ag.

FIG. 24 shows AFM characterization of these surfaces described inexample 4.

FIG. 25 shows optical spectra as a function of electrochemical potential(E) for a surface with high Au coverage.

FIG. 26 shows the changes in optical properties accompanying chemicaldeposition of Ag onto preformed Au colloid monolayers.

FIG. 27 shows the uv-vis optical properties of a surface coated withcolloidal metal particles that were stabilized with PEG.

FIG. 28 shows the optical spectrum of a substrate coated withpoly(allylamine) hydrochloride and immersed in a solution of colloidalAu.

FIG. 29 shows the optical spectrum of a colloid monolayer on SnO₂electrode;

FIGS. 30a-c show a tapping-mode AFM images of Au colloid multilayer filmpreparation on APTMS-derivatized glass using 12 nm Au particles,wherein;

FIG. 30a shows an image of a submonolayer;

FIG. 30b shows an image of a submonolayer after 2 exposures to2-mercaptoethanol/Au colloid; and

FIG. 30c shows an image of the submonolayer(bottom left) after 5exposures to 2-mercaptoethanol/Au colloid, images of FIGS. 30a-c eachbeing 1 mm×1 mm with 0-100 nm z-scale.

FIG. 31 shows Au particle coverage dependence (in particles/cm²) ofuv-vis/near-IR transmission spectra of Au colloid multilayers (preparedby successive, repeated immersion of a glass slide derivatized withAPTMS and a 12-nm diameter Au colloid monolayer into (a) 4 mM2-mercaptoethanol and (b) 17 nM, 12 nm-diameter Au);

FIG. 32 show the percent transmission at 1500 nm (+) and 2500 nm (O) asa function of Au colloid coverage for a Au colloid multilayer. Thesedata were obtained from FIG. 1 of the manuscript.

FIG. 33 shows a log plot of DC resistance versus Au particle coveragefor Au colloid multilayers. See FIG. 29 for details of samplepreparation.

FIG. 34 shows Au particle coverage (in particles/cm²) vs. the number ofimmersions into 12-nm colloidal Au in which one immersion corresponds toa monolayer.

FIG. 35 shows a diagram of a device for determining the presence of anantigen protein.

FIG. 36 shows the coupling of a colloid-based SERS with capillaryelectrophoresis.

FIG. 37 illustrates a technique for forming a self-assembled metalcolloid monolayer that exhibits continuous nanometer-scale gradient ofmixed-metal compositionand surface architecture. Such gradients areuseful for optimizing surface characteristics of the metal colloidmonolayer for a given application.

FIG. 38 shows UV-vis waves of 9 different Au/Ag coatings of a stepwisegradient combinatorial surface. The sample was submerged into Au in 3steps of 0.83 cm, with 50 minute pauses, turned 90°, and lowered into anAg⁺ solution in 3 steps of 0.67 cm, with 10 minute pauses resulting in amaximum Au exposure time of 2.5 hrs, and Ag exposure time of 30 min. Auand Ag exposure times for spectrum ω are analogous to the exposures forthe morphology represented by AFM image C in FIG. 42.

FIG. 39 shows SERS response of spatially discrete areas on a 2.5 cm×2.0cm continuous gradient combinatorial surface (4 hrs for Au, and 20minutes for Ag, respectively). Each box represents a SERS spectrumcollected in a 0.2 cm×0.2 cm area on the sample; each spectrum's x-axisis Raman shift (1300 cm⁻¹ to 1650 cm⁻¹) and y-axis is intensity (0counts per second (cps) to 35000 cps). The numbers located in each boxindicate the background subtracted intensity of the 1610 cm⁻¹ ringstretch mode of p-NDMA, and box locations in space correspond to theacquisition position on the sample; bold vertical lines indicate a 0.2cm jump in position. Particle coverage increases from left to right, andparticle size increases from top to bottom. Experimental conditions;[p-NDMA]=0.3 mM, 20 mW of 530.9 nm excitation, 1 cm⁻¹ step, 1-sintegration, 5 cm⁻¹ band pass.

FIG. 40 shows background subtracted SERS intensity map in counts persecond (cps) for the 1610 cm⁻¹ symmetric ring stretch of p-NDMA on alarge scale continuous gradient combinatorial surface, 0-12 hr for Au,0-30 min for Ag. Each box represents one SERS spectrum in a 0.1 cm×0.1cm area, total sample size is 2 cm×2 cm. Experimental conditions;[p-NDMA]=2.5 mM, 25 mW of 647.1 nm photons focused at the sample; 1 cm⁻¹step, 1 s integration, 5 cm⁻¹ band pass.

FIG. 41 shows background subtracted SERS intensity in counts per second(cps) for the 1168 cm⁻¹ phenyl-nitroso stretch of p-NDMA as a functionof sample position. Each shaded box represents one SERS spectrumcollected in a 0.1 cm×0.1 cm area. Regions A-D correspond to AFM imagesin FIG. 39. Experimental conditions: [p-NDMA]=2.5 mM; 30 mW of 647.1 nm,2 mm diameter spot at the sample; 1 cm⁻¹ step, 1 s integration, 5 cm⁻¹band pass.

FIG. 42 shows AFM images (500 nm×500 nm) of MPTMS-derivatized glassslides with varying coverages of 12-nm diameter colloidal Au and varyingquantities of deposited Ag. Images A-D correspond to regions delineatedin FIG. 38. Z-axis (vertical) gray scale: black=0 nm, white=35 nm.

FIG. 43 shows wavelength-dependent SERS maps from a single sample of aAu/Ag continuous gradient colloidal Au on3-mercaptopropylmethyl-dimethoxysilane (MPMDMS)/glass. Intensities havebeen normalized to 30 mW of 514 nm excitation. Each square represents 1spectrum in a 2 mm×2 mm area, with the peak intensities calculated fromthe 1610 cm⁻¹ band of trans-4,4′-bis-pyridylethylene (BPE). Experimentalconditions: 30 minutes Au exposure, 40-minute Ag exposure. [BPE]=10 mM;two collections of a 2-s integration, 5 cm⁻¹ band pass.

FIGS. 44A, 44B and 44C shows wavelength dependent SERS maps from threesamples of continuous combinatorial gradients based on 14- and 28-nmdiameter colloidal Au (Au colloid/MPMDMS/glass). Exposure times on theright-hand image (Au exposure=180 minutes, 5× diluted colloid, Agexposure=40 minutes) were based on the experimental findings for theleft-hand image (30 minutes Au exposure, 40-minute Ag exposure).Experimental conditions are the same as in FIG. 43.

FIG. 45 is a bar graph showing the number of particles per square micronas derived from analysis of AFM images (Au colloid/MPMDMS/glass).Numbers on the bars represent the mean particle height (in nm) for theimages as obtained via a section analysis.

FIG. 46A shows the optical spectra of 14-nm diameter colloidal Au cladwith Ag. Each spectrum has the same exposure time to Ag, but differentamounts of Au particles with Au₃>Au₂>Au₁. The samples comprisedmonolayers of Au colloid on MPMDMS-coated glass slides.

FIG. 46B shows the optical spectra of 14-nm colloidal Au clad with Ag.The spectrum of each of FIG. 46A and FIG. 46B has the same exposure timeto Au, but different durations of Ag exposure with Ag3>Ag2>Ag₁. Thesamples comprised monolayers of Au colloid on MPMDMS-coated glassslides.

FIGS. 47A and 47B show SERS spectra for the samples shown in FIG. 46Aand 46B, respectively. Experimental conditions: [BPE]=10 mM; 30 mW of647.1-nm excitation; two collections of a 2- s integration, 5 cm⁻¹ bandpass.

FIGS. 48A, 48B and 48C show AFM images (1 μm×1 μm, with 80 nm Z-scale)for the samples with varying Ag coverages described in FIGS. 46A and46B, and FIGS. 47A and 47B.

FIGS. 49A, 49B and 49C show AFM images (1 μm×1 μm, with 80 nm Z-scale)for the samples with varying Au coverages described in FIGS. 46A and46B, and FIGS. 47A and 47B.

FIG. 50A shows the optical spectra of Ag-clad, 28-nm diameter colloidalAu monolayers on MPMDMS/glass (Ag₃>Ag2>Ag₁).

FIG. 50B shows SERS spectra for Ag-clad, 28-nm diameter colloidal Aumonolayers on MPMDMS/glass. Experimental conditions for FIGS. 50A and50B:[BPE]=10 mM; 30 mW of 647.1-nm excitation; two collections of a 2- sintegration, 5 cm⁻¹ band pass.

FIGS. 51A, 51B and 51C show AFM images (1 μm×1 μm, with 80 nm Z-scale)for the samples described in FIGS. 50A and 50B.

DETAILED DESCRIPTION OF THE INVENTION

Below we describe data regarding the preparation, characterization, andapplications of self-metal colloid monolayers. Experimental examples areincluded.

Two lines of evidence demonstrate that immobilized particles are locatedsolely at the surface of, and not embedded within, the organic film. (i)Colloidal particles are very tightly attached to the polymer (whenstored in water, no particle dissociation occurs after 1 year), yetmonolayer formation does not occur on polymers with pendant methyl ormethoxy groups. These data indicate that multiple specific covalentinteractions between polymer functional groups (which are orientedtoward the solution) and the particle surface are necessary forimmobilization. (ii) Although SERS spectra for adsorbates from solutionare easily obtained (see below), the SERS spectra of organosilanepolymer films underneath Au monolayers are quite weak. This contrastswith published SERS studies of colloid/polymer mixtures [P. Matejka, B.Vlcková, J. Vohlidal, P. Pancoska, V. Baumruk, J. Phys. Chem. 96, 1361(1992); P. C. Lee and D. Meisel, Chem. Phys. Lett. 99, 262 (1983)], anddemonstrates that the surface of immobilized metal particles isaccessible to solvent. In accord with this finding is our observationthat the optical spectrum of Au colloid monolayers on transparentsubstrates depends on the dielectric constant of the surrounding medium.

The optical properties of colloidal Au and the nature of self assemblyoffer an unprecedented opportunity to monitor surface evolution in realtime. The time course of Au colloid monolayer formation on a glass slidecoated with polymerized 3-aminopropyltrimethoxysilane (APTMS) is shownin FIG. 2. Binding of 12-nm diameter Au particles to amine groups on thesurface is indicated by an absorbance feature at 520 nm, the location ofthe Mie resonance for isolated small Au particles. As the particlecoverage increases, interparticle spacing becomes small compared to theincident wavelength, and a new feature corresponding to a collectiveparticle surface plasmon oscillation grows in at ˜650 nm. This featureis responsible for the pronounced SERS activity of collections ofcolloidal Au particles. Accordingly, when a colloid monolayer in variousstages of formation is placed in a solution containing the adsorbatetrans-1,2-bis(4-pyridyl)ethylene (BPE), the SERS intensity for the ringstretch at 1610 cm⁻¹ closely tracks the magnitude of the absorbance at650 nm. Immersion time is one of four routes we have found to alter therate or extent of surface formation, the others being choice oforganosilane functional group (rate of surface formation forSH>NH₂>>CN), colloid concentration, and the presence or absence of anadsorbate on the colloidal particle.

FIG. 2A shows kinetics of Au colloid monolayer formation byultraviolet-visible (uv-vis). A series of uv-vis spectra of Aucolloid-functionalized glass slides in H₂O obtained with an HP-8452Aspectrophotometer. Cleaned (4:1 H₂SO₄:H₂O₂, 70° C.) rectangular glassslides ( 0.9 mm×25 mm) were placed into a dilute solution of3-aminopropyltrimethoxysilane (APTMS) (0.3 ml: 3 ml of CH₃OH) for 12hours and rinsed with CH₃OH upon removal. The polymer-coated slides werethen immersed in a 17 nM solution of 12-nm diameter colloidal Auparticles (wavelength maximum=520 nm) [Garrell, R. L. Anal. Chem. 1989,61, 401A-11A; Tran, C. D. Anal. Chem. 1984, 56, 824-67,20]. At each timeindicated (and at several others not shown), the slide was removed fromthe Au colloid solution, and an optical spectrum was recorded in H₂O,followed by a SERS spectrum in 4 mM BPE in 95:5 H₂O:CH₃OH (20 mW 632.8nm, Spex 1403 double monochromator, Hamamatsu R928 photomultiplier tube,bandpass=7 cm⁻¹, scan rate=1 cm⁻¹s⁻¹, integration time=1 s). FIG. 2Bshows kinetics of Au colloid monolayer formation by SERS. SERS intensityfor the 1610 cm⁻¹ band versus absorbance at 650 nm. Other bands in theBPE SERS spectrum evolve with identical kinetics.

This high degree of control over surface formation has importantramifications for reproducibility, a long-standing complication in SERSresearch. For example, when BPE was adsorbed to eight identical Agcolloid monolayers on glass, the greatest variation in integrated peakintensity for the 1610 cm⁻¹ band was less than 8%. Similarly, for fivedifferent locations on a single substrate, the greatest difference wasonly 5%. As these values incorporate intrinsic errors associated withvariation in laser power and sample positioning, the actual samplereproducibility is significantly better. This reproducibility extends tothe nanometer scale, where Au and Ag colloid monolayers have been imagedusing transmission electron microscopy (TEM), field emission scanningelectron microscopy (FE-SEM), and atomic force microscopy (AFM). Arepresentative TEM image of an Au colloid monolayer prepared on anSiO_(x)-coated Formvar surface is shown in FIG. 3. The Au particles areconfined to a single layer, and the vast majority of particles areisolated from each other, unlike previous TEM studies of SERS-active Auand Ag colloids. Furthermore, the large field of view available with TEMallows us to conclude that particle aggregation has been eliminated overthe entire sample. Similar conclusions obtain from large-field FE-SEMimages and from multisite tapping-mode AFM images of Au-modified glasssurfaces. The AFM image from a glass slide coated with3-aminopropylmethyldimethoxysilane indicates a roughness of 1 to 3 nm,notwithstanding a few isolated locations where the polymer roughnessapproaches 8 to 10 nm. This roughness scale is typical for organosilanefilms on glass or quartz. Immobilization of 12-nm colloidal Au particlesto a coverage equivalent to that shown for 180 to 210 min in FIG. 2yields a surface with features 12 to 20 nm high and 20 to 30 nm wide.The increased dispersion in particle size relative to TEM results fromconvolution of the true particle size with the AFM tip size, but arenevertheless of sufficient quality to conclude that the surface iscomposed of a monolayer of separated particles, in agreement with FE-SEMimages on similar substrates. Importantly, we have demonstrated that thespacing obtained on these colloid-based surfaces is sufficient to yieldSERS enhancement. The bottom panel of FIG. 3B shows the SERS spectrum ofBPE adsorbed onto the derivatized TEM grid pictured in the top panel.For comparison, the Raman scattering spectrum of an equivalent amount ofBPE deposited onto an unmodified SiO_(x)-coated TEM grid is also shown.The intensity difference in these two samples clearly demonstrates theenhancing properties of colloid-based surfaces.

FIG. 3A shows an image from a Formvar-coated Cu TEM grid which had beensputter-coated with a thin layer of SiO_(x) (Ted Pella, Inc.), treatedfor 2.5 hours in neat 3-cyanopropyldimethylmethoxysilane, rinsedexhaustively with CH₃OH upon removal, and immersed for 12 hours incolloidal Au (12 nm diameter) [Garrell, R. L. Anal. Chem. 1989, 61,401A-11A; Tran, C. D. Anal. Chem. 1984, 56, 824-6]. Imaging wasperformed on a JEOL 1200 EXII instrument operated at 80 kV acceleratingvoltage. The area depicted is 0.28 mm² and is representative of thesample surface. FIG. 3B shows a SERS spectrum (upper) of 5 ml of 1 mMBPE drop-coated onto the surface of the derivatized TEM grid (100 mW,647.1 nm, 5 cm⁻¹ bandpass, 2 cm⁻¹ step, 2 s integration). Forcomparison, an identical quantity of BPE was drop-coated onto anunderivatized SiO_(x) grid; the Raman spectrum from this sample is shown(1 cm⁻¹ step, 1 s integration).

Another important feature of film-supported metal colloid monolayers isthat the particles are subject to electrochemical potentials applied tounderlying conductive substrates. Consequently, like SERS-activeelectrodes, Ag colloids immobilized on Pt exhibit an electrochemicalpotential-dependent SERS intensity for adsorbed pyridine (FIG. 4).Identical maximas for the two surfaces in the intensity versus potentialplots suggests that the voltage drop across the polymer film is minimal.Voltammetry at colloid-based surfaces also resembles that at macroscopicelectrodes. The first reduction wave for methyl viologen (MV²⁺) ismarkedly rectified at an organosilane-coated Pt electrode (FIG. 4,inset) but returns upon immobilization of Au particles. The slightlybroadened peak-to-peak separation is expected for an array of closelyspaced microelectrodes. Considering the demonstrated biocompatibility of5 to 20 nm diameter Au particles, the ability to make electrochemicalmeasurements at Au colloid monolayers suggests possible electrode-basedbiosensor applications.

FIG. 4 shows an electrochemical potential dependence of SERS intensityof the 1006 cm⁻¹ band of pyridine on an Ag colloid monolayer on Pt andon bulk Ag [Albrecht, M. G.; Creighton, J. A. J. Am. Chem. Soc. 1977,99, 5215-7]. The monolayer was prepared as follows: Clean Pt foil wasplaced into neat APTMS for 4 hours. After rinsing with triply distilledH₂O and air-drying, the polymer-coated foil was dipped in Ag colloidsolution [Soper, S. A.; Ratzhlaff, K. L.; Kuwana, T. Anal. Chem. 1990,62, 1438-44] for 1 hour. The derivatized foil was then rinsed withtriply distilled H₂O and air-dried. In the absence of colloidal Ag, nopyridine SERS spectra were observed at any potential. See FIG. 2 forspectral acquisition parameters. Inset: Voltammograms (100 mV/s, N₂atmosphere) of 5 mM MV²⁺ in 0.1 M Na₂SO₄ on three surfaces: unmodifiedPt, Pt coated with surface-polymerized3-mercaptopropylmethyl-dimethoxysilane (MPMDMS), and Pt coated withMPMDMS and derivatized with 15-nm diameter Au particles (5 hours in neatsilane, rinsed, 4 hours in colloidal Au).

Interparticle spacing in preformed Au monolayers can be further reducedby chemical deposition of an Ag coating; increased interparticlecoupling because of decreased spacing and concomitant changes indielectric properties lead to a dramatic increase in SERS activity. Theoptical and SERS spectra before and after deposition of Ag onto 18 nmdiameter colloidal Au are shown in FIG. 5. Initially, relatively largeinterparticle spacing is indicated by the absence of a collectiveparticle surface plasmon band in the ultraviolet-visible and by a weaklyenhanced SERS spectrum for adsorbed para-nitrosodimethylaniline(p-NDMA). Silver deposition causes a large increase in extinction at allwavelengths as well as a shift in λ_(max) from 520 to 386 nm. The shiftin energy of and increased extinction at λ_(max) concur withexpectations based on a computer algorithm for predicting the opticalproperties of isolated coated particles [C. Bohren and D. R. Huffman,Absorption and Scattering of Light by Small Particles (Wiley, New York,1983)]; best agreement between the experimental and model data isreached with a 4-nm Ag coat (to make 26-nm diameter particles) [Theoptical constants for Au and Ag were taken from R. H. Morriss and L. F.Collins, J. Chem. Phys. 41, 3357 (1961). These values were fit toexponential curves to generate continuous values between 300 and 700nm]. The exceptional SERS activity (enhancement factor=105) [anenhancement factor (EF) of 5.7×10⁵ was calculated for the Ag-coatedsurface by comparing the ratios of background-corrected intensities fora SERS spectrum and a solution spectrum in units of counts s⁻¹ watt⁻¹molecule⁻¹, and averaging the EF values obtained for six differentcommon peaks—low signal/noise precluded calculation of accurate EFs forthe as prepared Au sample] of these substrates reflects optimization ofthe Ag coating thickness for this particular particle size and spacingof colloidal Au—even greater enhancements may be possible with othercombinations.

FIG. 5 shows the effect of Ag coating on the uv-vis and SERS spectra ofpreformed Au colloid monolayers. The initial substrates were prepared asin FIG. 2, except that the organic film was formed from reaction with2-(trimethoxysilyl)ethyl-2-pyridine (PETMS) for 24 hours. Silver coatingwas performed by immersing Au colloid monolayers into a 1:1 mixture ofLI Silver enhancer and initiator solutions (Nanoprobes Inc., StonyBrook, N.Y.) for 13 min. The SERS spectra were of 0.5 mM p-NDMAsolutions in CH₃OH. Optical spectra (inset) were measured in H₂O.Instrumental parameters were described in FIG. 2. When Ag is depositedfrom the plating solution onto a PETMS-derived polymer on glass in theabsence of colloidal Au, no SERS intensity could be observed for thesame p-NDMA solution, irrespective of coating time.

More detailed characterization of these surfaces follows. The top panelof FIG. 6 shows optical spectra for solutions of isolated and aggregated13-nm diameter colloidal Au particles in H₂O. The unaggregated sol,which has a particle concentration of 17 nM, has a λ_(max) of 520 nm.The physical nature of this surface plasmon mode, which gives colloidalAu its characteristic intense burgundy color, is well-understood, as areits dependence on particle size and shape. When the interparticlerepulsive forces are sufficiently screened by molecular adsorption,irreversible aggregation occurs and generates a new red-shifted featurein the optical spectrum centered between 600-800 nm. The intensity andλ_(max) of this feature scale with the extent of aggregation, with largeaggregates exhibiting increased extinction and red-shifted peaks. This“aggregated” band results from coupling of surface plasmons betweenclosely-spaced particles. It has been amply demonstrated, boththeoretically and experimentally, that the SERS-activity of aggregatedcolloidal Au arises from this interparticle coupling. In aggregatedsols, the particles are physically connected, but it is important tonote that direct contact is not needed to observe collective plasmonmodes: as long as the spacing between particles is small compared to thewavelength of light, these collective plasmon modes can be observed.Uv-vis is thus particularly well-suited for analyzing our samples, sincethe optical spectra of Au colloid monolayers on transparent substratesis easily measured. The same cannot be said of most SERS substrates witha notable exception being those prepared by Roark et al. [Roark, S. E.;Rowlen, K. L. Appl. Spectrosc. 1992, 46, 1759-61; Roark, Shane E.;Rowlen, K. L. Chem. Phys. Lett. 1993, 212, 50; Roark, Shane E.; Rowlen,K. L. Anal. Chem. 1994, 66, 261-70]. Moreover, colloid self-assemblyprovides a means of tuning surface optical properties through control ofinterparticle spacings.

The diversity of optical properties attainable through self-assembly ofcolloidal Au is illustrated in the bottom panel of FIG. 6. Use of twodifferent organosilanes and two sizes of colloidal Au particles yieldsfour distinct surfaces, as evidenced by different optical spectra. Incomparison to the data in the top panel, it is clear that interparticlecoupling is not as pronounced as for aggregated colloidal solutions.Using 13-nm diameter particles at 0.15 monolayer coverage, there areroughly 1×10¹¹ particles in a 1 cm² monolayer which, using a 15-nm slabthickness, are in a volume of 15×10⁻¹⁰ liters. This translates to asurface concentration of 1×10⁻⁴ M, versus 17 nM in solution. Despitethis 4 order-of-magnitude increase in concentration (one that cannot bemaintained in solution without aggregation), the particles remaindistinct; this lack of surface aggregation is additional strong evidencefor specific interactions between the surface of Au and the polymerfunctional groups and the high stability suggests that multiple linkagesmust be present. These data are reinforced by the absence of Au or Agimmobilization on polymers derived from trimethoxypropylsilane, whichlacks a high-affinity functional group. The key point is that thepolymer-particle interaction, an adjustable parameter, controls theparticle density, which in turn dictates the optical properties.

FIG. 6A shows absorbance spectra for solutions of Au colloid (13 nmdiameter). Unaggregated Au has a λ_(max) at 520 nm, while aggregated Auexhibits a second, red-shifted absorbance centered at 700 nm. The solwas aggregated via addition of a small volume of concentrated NaClsolution. FIG. 6B shows absorbance spectra of quartz slides derivatizedfor 14 h in neat silane and for 24 h in colloidal Au: (A) APTMS, 30 nmAu; (B) APTMS, 13 nm Au; (C) MPMDMS, 30 nm Au; (D) MPMDMS, 13 nm Au.

Thus, for surfaces with the same polymer and containing a single size ofparticle, difference in optical properties must be attributed todifferences in coverage (and therefore, average interparticle spacing).For example, spectra B and D are of immobilized 13 nm-diameter Au onquartz substrates derivatized with amino (APTMS)- and sulfhydryl(MPMDMS)-functionalized siloxane polymers, respectively. Relativelystrong interparticle coupling is found in B, as evidenced by thepresence of the collective surface plasmon absorbance feature, but isabsent (or significantly weaker) in D. Since the area probed by theuv-vis beam is constant, and since the Au particle coating is homogenousover the entire surface, the stronger interparticle coupling resultsfrom an increased particle density. Whether this difference isattributable to a higher concentration of pendant functional groups insurface-confined APTMS than for MPMDMS, an increased affinity of Au foramine over sulfhydryl, or some other factor is under investigation.

These surfaces are fundamentally different from those prepared byevaporation of drop-coated colloidal Au solutions. Evaporated substratesexhibit complete colloid aggregation, sometimes to the extent ofproducing films that to the eye look like bulk Au. In contrast, theprotocol described herein involves no bulk aggregation on the surface.Furthermore, with adequate rinsing between the polymer formation andcolloid derivatization steps, there is no aggregation of particles insolution; immersion of the polymer-functionalized substrate into acolloidal Au solution, and subsequent removal of the colloid-derivatizedsurface, does not appreciably change the optical spectrum of thecolloidal Au solution. We have also shown that colloid immobilization isnot a sedimentation reaction by performing derivatizations upside down.Thus, immobilization of colloids on a polymer-functionalized glasssubstrate suspended upside down in solution yields colloidal surfacesindistinguishable from those obtained by complete immersion. Similarly,polymer-coated TEM grids can be derivatized with Au by flotation onaqueous colloidal solutions.

Once attached, the binding of colloidal Au to derivatized surfaces isextremely strong and essentially irreversible. There is very littlechange in the optical spectrum of an Au colloid-based monolayer afterstorgage for 7 months in H₂O (FIG. 7). Ag-based surfaces are also verydurable, with no loss of particles over 2 years. For Ag, a shift ofλ_(max) from 396 to 420 nm may reflect particle aging, as has previouslybeen shown for Ag colloids [Henglein, A. J. Phys. Chem. 1993, 97,5457-71 and references therein. An alternative explanation for the shiftis that bacterial growth in solution, against which no precautions weretaken, leads to adsorption of protein on the particle surface (see FIG.3)]. More importantly, these surfaces are rugged enough to surviveexposure to appreciable concentrations of aggregating agents. Thus,exposure of an MPMDMS-based Au colloid surface to 5 mM mercaptoethanoldoes not alter the optical spectrum, indicating that particleaggregation has not taken place (data not shown). It is significant thatthe same concentration of aqueous mercaptoethanol instantaneouslyaggregates colloidal Au and Ag in solution. The high durability of thesesubstrates is further manifested by their resistance toward ligandexchange: solution RS⁻does not displace surface RS⁻/Au bonds. Indeed,neat mercaptoethanol is needed to effect particle removal. Likewise,immersion of an Au-coated substrate into a solution of H₂O at 75° C. fora period of one hour had no effect on the optical spectrum. Equivalentstabilities are found for surfaces based on NH₂-Au linkages.

FIG. 7A shows absorbance spectra for a glass slide derivatized for 24 hin APTMS (diluted 1:4 with CH₃OH) and for 11 days in colloidal Au (13nm). FIG. 7B shows absorbance spectra for a glass slide derivatized for14 h in neat MPMDMS and for 1 week in colloidal Ag. After the initialoptical spectra were recorded, slides were stored in H₂O until the finalspectra were taken.

Surfaces based on non-covalent interactions (FIG. 1) possess opticalproperties completely analogous to those prepared by covalent attachment(FIG. 8). An important aspect of these data and of the concept describedin FIG. 1 is that the biological activity (i.e. biotin binding) ofstreptavidin adsorbed on colloidal Au is necessarily retained:unmodified Au particles and particles coated completely with a proteinthat doesn't specifically bind biotin (i.e. BSA) do not lead to surfaceformation in the presence of biotinylated substrates. The retention ofbiological function contrasts sharply with streptavidin adsorbed at bulkAu surfaces, for which biological activity is compromised [Ebersole, R.C.; Miller, J. A.; Moran, J. R.; Ward, M. D. J. Am. Chem. Soc. 1990,112, 3239-41]. The use of colloidal Au as a histochemical andcytochemical marker is based on the tendency of proteins adsorbed tosmall Au particles to retain their biological function. A majoradvantages of these surfaces, then, is their biocompatibility. Becausethey are composed of isolated colloidal particles, the behavior of thesurface mirrors the behavior of particles in solution. The creation ofmacroscopic metal surfaces with high, nanometer-scale biocompatibilityis important for biosensor applications, and reinforces the importanceof maintaining some interparticle spacing, for only under theseconditions can single particle behavior toward biomolecules be assured.Of course, the biomolecule itself may be the spacer . A final comment onFIG. 1 concerns the use of a coating protein to completely isolate Auparticles. For the data in FIG. 8, BSA was used, meaning that eachparticle had multiple BSA molecules adsorbed for each streptavidinbound. (Note the peak shift of roughly 20 nm for λ_(max), reflecting achange in local dielectric constant of proteins relative to H₂O.)However, this choice is arbitrary; it is possible to prepareparticle-based Au surfaces where each particle is pre-coated with aprotein of interest.

Transmission Electron Microscopy

Direct evidence concerning the morphology and interparticle spacingcomes from transmission electron microscopy studies of colloidal Aubound to polymers on TEM grids. These were prepared using commerciallyavailable formvar-coated Cu TEM grids possessing a thin sputter-coatedoverlayer of SiO₁. Careful treatment of these fragile surfaces withorganosilane followed by colloidal Au yielded surfaces that could bedirectly imaged. FIG. 9A shows two magnifications of a surfacederivatized in this manner with 13-nm colloidal Au. The areas shown inthese micrographs are roughly 4.0 mm² for the top panel and 0.2 mm² forthe bottom panel, and are representative of the entire sample. FIGS. 9,9A & 9B show TEM micrographs of a colloidal Au surface prepared byderivatizing an SiO_(x)-coated TEM grid for 2.5 h in neat CPDMMS and for12 h in 13 nm Au colloid. Areas depicted are approximately as follows:(top) 4.0 mm², (bottom) 0.2 mm².

Examination of these images verifies several critical aspects of thestrategy delineated by FIG. 1: (1) there is a single two-dimensionalsubmonolayer of colloidal Au; (2) the particles are closely spaced butnot aggregated in two dimensions; (3) the particle coverage is uniformover areas macroscopic compared to the particle size; (4) the roughnessis uniform and defined solely by the particle diameter; and (5) thereappears to be a limitation to the number of particles that can be boundper unit area, with only 15%-20% of the surface covered. The observeddistribution of particles extends over macroscopic areas, i.e. 3 mm×3mm, the size of the TEM samples that we typically prepare.

It is well-known that aggregation of colloidal Au produces fractalclusters and strings. In the Creighton group's groundbreaking work oncolloid SERS, two-dimensional strings of SERS-active particles wereimaged by TEM [Jeanmaire, D. L.; Van Duyne, R. P. J. Electroanal. Chem.1977, 84, 1-20]. Such species are not seen here, and the smallpercentage of dimers and trimers are invariably found in colloidal Ausolutions as prepared. Weitz and co-workers have characterized thefractal dimension of aggregated Au and correlated it to SERS activity[Weitz, D. A.; Lin, M. Y. Surf. Sci. 1985, 158, 147-64]. Again, theselarge aggregates are not seen in images of carefully prepared surfaces.By way of contrast, FIG. 9C depicts a TEM image of colloidal Au on anon-functionalized, formvar-coated grid. Three-dimensional clusters ofparticles are clearly present in addition to isolated particles; theinability to achieve a good focus further signifies the existence ofmultiple layers of colloidal particles. Such species are not observed onthe grids from which the data in FIGS. 9A and 9B was extracted. Rather,observation of closely-spaced, predominantly unaggregated colloidalparticles confirms the arguments made above based on opticalspectroscopy. The fact that all the colloidal particles are confined tonearly a single plane, as evidenced by good focus over large areas,suggests that for these surfaces, the roughness of the underlyingorganosilane film and/or substrate is comparable to the particlediameter or smaller. In accord with this notion, several studies oforganosilane polymer films on smooth surfaces indicate a thickness <20 Å[Dressick, W. J.; Dulcey, C. S.; Georger, J. H., Jr.; Calabrese, G. S.;Calvert, J. M. J. Electrochem. Soc. 1994, 141, 210-20; Karrasch, S.;Dolder, M.; Schabert, F.; Ramsden, J.; Engel, A. Biophys. J. 1993, 65,2437-46; Nakagawa, T.; Ogawa, K.; Kurumizawa, T. Langmuir 1994,10,525-9].

The tendency toward even spacing between particles observed in FIG. 8results from electrostatic factors. It is known that colloidal particlesare negatively charged and thus naturally repel one another, and thataggregation occurs only under conditions where this interparticlerepulsion is screened. Within this framework, the protocol describedhere is self-assembly, in that long range order arises from secondaryinteractions between individual particles, as opposed toparticle-surface interactions.

Surface Enhanced Raman Scattering (SERS)

One of the principle objectives of assembly of macroscopic metalsurfaces exhibiting controlled roughness is to prepare well-defined,reproducible SERS-active substrates. The optical spectra show that theparticle spacing is small compared to 1, suggesting that these particlearrays should be SERS-active. FIG. 10A shows the SERS spectrum of 5 nmolof BPE drop-coated onto a colloid monolayer on a functionalized TEMgrid, as well as the SERS spectrum of the adsorbate-free surface. Thesedata are extremely significant because they were obtained on the sametype of surface imaged by TEM in FIG. 8. In the absence of adsorbate, nomajor features are observed in the Raman spectrum, indicating SERS fromthe polymer underlayer is weak. Typically, a low energy mode is observedfor the S-Au vibration from MPMDMS-derived films, but little else iseasily discerned.

FIG. 10A shows SERS spectrum of 5 ml of 1 mM BPE drop-coated onto anSiO_(x) coated TEM grid derivatized with MPMDMS and 10 nm colloidal Au.Excitation source: 647.1 nm, 100 mW; 2 cm⁻¹ step, 2 s integration; 5cm⁻¹ bandpass. The bottom spectrum was taken of the substrate surfaceprior to adsorption of BPE. The surface topography was identical to thatdepicted in FIG. 2. FIG. 10B shows SERS spectrum of BPE (10 ml, 1 mM inCH₃OH) drop-coated onto a colloidal Au (10 nm diameter) monolayerprepared on a HO₂C(CH₂)₁₈SH/Au/Cr/Si substrate (upper). Prior toadsorption of BPE, a background spectrum of the colloidal Au substratewas run (lower). Excitation source: 647.1 nm; 150 mW (BPE), 100 mW(background); 2 cm⁻¹ step, 1 s integration; 5 cm⁻¹ bandpass.

It is important to understand the factors responsible for the observedSERS behavior of these substrates: why do we see BPE and not the polymerunderlayer? Most SERS studies of polymers show a number ofpolymer-related bands. However, in those studies, the polymerscompletely surrounded the colloidal particles, while in this work, ifthe idealized geometry in Scheme I is reasonable (as the TEM data inFIG. 8 suggest is the case), only a small fraction of the colloidsurface contacts polymer. For a particle with diameter=2r, the fractionof a sphere's surface area covered by a polymer with x nm verticalflexibility (over a horizontal distance 2r) is equal to: $\begin{matrix}\frac{\cos^{- 1}\left\lbrack {\left( {r - x} \right)/r} \right\rbrack}{p} & (1)\end{matrix}$

In our system, using r=6 nm and x=1 nm (a vast overestimate, consideringthe length of the —CH₂CH₂CH₂A tail is itself <1 nm), the fraction oftotal surface area exposed to polymer is 0.186. In other words, theratio of adsorbate molecules at monolayer coverage to polymer tails isat minimum 5:1, and more likely closer to 10:1.

Another factor influencing the SERS enhancement is the Raman scatteringcross-section. BPE is an exceptionally strong scatterer, while alkanesyield very weak Raman spectra. Bryant et al. have measured SERS spectraof octadecanthiols on Au foil; using a CCD, very long (10 min)integration times were required. A third consideration is the magnitudethe electric fields responsible for the electromagnetic enhancement. Thelargest fields are expected to occur in the plane of the particles, notin the plane normal to the substrate, i.e. Raman spectra of moleculesadsorbed in this region are enhanced to a greater extent than thoseadsorbed elsewhere, and from our calculations above, only BPE can accessthis region. Finally, chemical enhancement effects in SERS certainlyfavor observation of enhanced Raman scattering from thenitrogen-containing BPE versus an alkane. The combination of thesefactors all favor observation of BPE SERS, and help explain theconsistently observed finding that, over the region between 400-1700cm⁻¹, very weak or no SERS spectra are seen for underlying films. Abenefit of these substrates is thus the lack of background spectra,simplifying the data acquisition process. On the other hand, thesesubstrates, like many others previously described, may not besufficiently enhancing to measure Raman spectra for weak scatterers orpoor adsorbates.

Because the Raman intensity of BPE adsorbed onto organosilane-coatedglass slides is too small for us to measure, we crudely estimated howenhancing these surfaces are by comparing the solution concentration [x]of BPE needed to yield the same normal Raman spectrum as we obtained fora BPE concentration [y] in the presence of an Au colloid monolayer.Typically, x/y≧10⁴. This number is in line with enhancements measured atroughened Au electrodes. The spectra yielding these enhancement factors(data not shown) are less than a factor of ten more intense than thedata in FIG. 10A. Thus, enhancement factors of >1000 can routinely beobtained from arrays of closely spaced but non-contacting particles.Importantly, identical spectra are obtained on substrates in which BPEwas adsorbed from solution; in fact, our experiments are routinelycarried out in this fashion.

Below are experimental details associated with colloid monolayerpreparation:

Materials. The following materials were obtained from Aldrich:HAuCl₄.3H₂O, AgNO₃, trisodium citrate dihydrate, trans-1,2bis(4-pyridyl)ethylene (BPE), and trimethoxy-propylsilane. The followingorganosilanes were obtained from Hüls America, Inc., and used asreceived: (3-aminopropyl)trimethoxysilane (APTMS),(3-cyanopropyldimethyl)methoxysilane (CPDMMS),(3-mercaptopropylmethyl)dimethoxysilane (MPMDMS), and3-cyanopropyltriethoxysilane (CPTES). Concentrated HCl, HNO₃, and H₂SO₄were purchased from J. T. Baker Inc., and 30% H₂O₂ was obtained fromVWR. CH₃OH (spectrophotometric grade) was obtained from EM Science; allH₂O was 18 MW, distilled through a Barnstead Nanopure water purificationsystem. Streptavidin, bovine serum albumin (BSA), and3-(N-maleimidopropionyl)biocytin (MP-biocytin) were purchased fromSigma. BPE was recrystallized several times from a mixture of H₂O andCH₃OH; the other materials were used as received. Substrates wereobtained as follows: glass and quartz microscope slides from FisherScientific and Technical Glass Products, respectively; SiO_(x)-coatedTEM grids from Ted Pella, Inc.; and self-assembled monolayers (SAMs) ofHS(CH₂)₁₈CO₂H on Au foil from literature procedures [Bain, C. D.;Troughton, E. B.; Tao, Y.-T.; Evall, J.; Whitesides, G. M.; Nuzzo, R. G.J. Am. Chem. Soc. 1989, 111, 321-35].

Colloid Preparation. All glassware used in these preparations wasthoroughly cleaned in aqua regia (3 parts HCl, 1 part HNO₃), rinsed intriply-distilled H₂O, and oven-dried prior to use. Au colloids wereprepared according to Frens [Frens, G. Nature Phys. Sci. 1973, 241,20-2] or Sutherland [Sutherland, W. S.; Winefordner, J. D. J. Colloid.Interface Sci. 1992, 48, 12941] with slight modifications. The followingstock solutions were prepared from triply-distilled H₂O that had beenfiltered through a 0.8 mm membrane filter (Gelman Scientific): 1%HAuCl₄, 38.8 mM sodium citrate, and 1% sodium citrate. Other solutionswere made fresh as needed using triply-distilled, filtered H₂O. Twotypical Au preparations and one Ag preparation are described below.

Preparation I: Using a 1 L round bottom flask equipped with a condenser,500 ml of 1 mM HAuCl₄ was brought to a rolling boil with vigorousstirring. Rapid addition of 50 ml of 38.8 mM sodium citrate to thevortex of the solution resulted in a color change from pale yellow toburgundy. Boiling was continued for 10 minutes; the heating mantle wasthen removed, and stirring was continued for an additional 15 minutes.After the solution reached room temperature, it was filtered through a0.8 mm Gelman Membrane filter. The resulting solution of colloidalparticles was characterized by an absorption maximum at 520 nm.Transmission electron microscopy (TEM) indicated a particle size of 13nm±1.7 nm (100 particles sampled). Preparation II: In a 1 L round bottomflask equipped with a condenser, 500 ml of 0.01% HAuCl₄ was brought to aboil with vigorous stirring. To this solution was added 7.5 ml of 1%sodium citrate. The solution turned blue within 25 s; the final colorchange to red-violet occurred 70 s later. Boiling continued for anadditional 10 min., the heating source was removed, and the colloid wasstirred for another 15 min. TEM data indicated an average diameter of 18nm±4.6 nm (89 particles sampled). Particle diameter was varied by addinglarger or smaller amounts of sodium citrate to decrease or increase theparticle size.

Ag colloid was prepared according to Lee and Meisel [Lee, P. C.; Meisel,D. J. Phys. Chem. 1982, 86, 3391-3395]. Using a heating plate and a 1 Lflask, a solution of 90 mg AgNO3 in 500 ml of triply distilled H₂O wasbrought to boiling with rapid stirring. To this solution was added 10 mlof 1% sodium citrate. Boiling continued for 30 min, after which time theflask was removed from the heat source, and the solution was dilutedwith triply distilled H₂O to obtain a final volume of 420 ml.

All colloids were stored at room temperature in dark bottles and weregenerally used within 1-2 months after preparation. Samples for particlesizing by TEM were prepared by drop coating 10 ml of the colloid onto aformvar-coated Cu grid and allowing the sample to dry. Average sizeswere determined by measuring diameters along a consistent axisthroughout the sample.

Protein-Colloid Conjugates. Streptavidin-labelled Au particles wereprepared using modifications of literature protocols [Liesi, P.; Julien,J.-P.; Vilja, P.; Grosveld, F.; Rechanrdt, L. J. Histochem. Cytochem.1986, 34, 923]. To 25 ml of colloidal Au (preparation I) were added0.725 ml of streptavidin (0.34 mg/ml in triply distilled H₂O) and 0.241ml of BSA (7.24 mg/ml in triply distilled H₂O). The protein-Auconjugates were observed to sediment within 24 hours.

Surface Derivatization. Substrates were cleaned prior to derivatizationas follows: glass and quartz, cut to dimensions of approximately 2cm×0.7 cm, were cleaned for 10 minutes in a bath consisting of 4 partsH₂SO₄ to 1 part 30% H₂O₂ at 60° C. The samples were rinsed inspectrophotometric grade CH₃OH and stored in this solvent until needed.SiO_(x)-coated TEM grids were cleaned in an ozone plasma for 30 minusing a home-built instrument. Cleaning often preceded use of the gridsby several weeks; during this period, the grids were stored in TEM gridholders in air.

Derivatization of glass and quartz substrates with alkoxysilanes wasaccomplished in the following manner: Clean substrates were submergedinto vials of silane diluted 1 part to 4 parts with spectrophotometricgrade CH₃OH. After a period of 24 h, the substrates were removed andrinsed profusely with CH₃OH to remove unbound monomer from the surface.At this point, silanized substrates were stored in CH₃OH until needed.Prior to derivatization with colloidal Au, the substrates were rinsedwith H₂O; they were then immersed in vials of colloidal Au for 24 h. Afinal H₂O rinse concluded the derivatization process. Similarly,carboxyl-terminated SAMs prepared on Au-coated silicon substrates wereimmersed in colloidal Au solutions for several days. The substrates werestored in H₂O until needed for analysis.

Due to their inherent fragility and small size, greater care wasrequired for the derivatization of TEM grids. Specifically, theSiO_(x)-coated TEM grids were immersed in neat silane for 3 h, followedby extensive methanol rinses and a H₂O rinse. The rinsing wasaccomplished by pipetting solvent across the grid surface, or byswishing the grid back and forth in a vial of solvent. Effort was madeto minimize the solvent flow perpendicular to the grid face in order tobetter preserve the formvar film. Finally, the grids were floated on acolloid solution for 12 h. Samples were rinsed with H₂O and allowed toair dry on filter paper prior to analysis.

Sample Preparation. Two methods were employed for mounting thesubstrates for SERS detection. The first method involved mounting thesubstrate via double-sided tape to a black mount positioned in the laserbeam (TEM grids, SAM substrate). In the second, the substrate (glass orquartz) was supported in the front of a quartz cuvette by means of ateflon block whose height was only ⅓ that of the sample. This cuvettecould be filled with solvent or empty. The cuvette rested in a snug,home-built cuvette holder. Both sample configurations were mounted on astage such that the sample position could be adjusted in all threedimensions. For measurements carried out in air, solutions of BPE inCH₃OH were drop-coated onto the substrate surface and allowed toevaporate; alternatively, the cuvettes were placed in cuvettescontaining known concentrations of BPE.

Instrumentation. SERS spectra were obtained with a Coherent Kr⁺ ionlaser, model 3000K, operated at 647.1 nm in TEM₀₀. Spectral scanning anddetection were accomplished through the use of a Spex Model 1404scanning double monochromator with a pair of 1800 grooves/mm gratingsand a thermoelectrically-cooled Hamamatsu R928 photomultiplier tubehoused in a Products for Research casing. Monochromator entrance andexit slits were typically set at 700 mm, and center slits were set at1400 mm to yield an effective band pass of 5 cm⁻¹. Grating movement andspectral acquisition were controlled using the DM3000 software providedby Spex. Plasma lines were filtered out of the incident laser beamthrough the use of a band pass filter (Ealing ElectroOptics) or apre-monochromator tuned to the 647 nm line (Optometrics). The laser beamwas focused onto the substrate sample at an angle of <30° from thesurface normal. Scattered radiation was collected and collimated with aMinolta 50 mm camera lens (f#1.2) and focused through a polarizationscrambler (Spex) onto the entrance slits of the monochromator.

Absorption spectra were obtained using a Hewlett-Packard 8452A diodearray spectrophotometer (2 nm spectral resolution, 1 s integrationtime). Again, substrates in quartz cuvettes were maintained in anupright position through the use of a teflon block. Transmissionelectron microscopy was performed on a JEOL Model 1200 EXII instrumentoperating at 80 kV accelerating voltage; the images were notmanipulated, altered, or enhanced in any way.

Below are eight (8) further examples of colloid monolayer experimentalprotocols:

EXAMPLE 1 Surfaces Made From Seeded Colloidal Au Particles

Glass slides (2.5 cm×0.8 cm×1 mm) were cleaned in a mixture of HCl:HN0₃(3:1). Slides were rinsed in H₂O and CH₃OH prior to derivatization for18 h in a solution of aminopropyltrimethoxysilane (diluted 1:5 inCH₃OH). The derivatized surfaces were rinsed extensively in CH₃OH andH₂O prior to immersion in solutions of colloidal are described below.After 24 hours, the colloid derivatization was complete.

Au nuclei (“seeds”) were prepared by adding 1 ml of 1% Na₃ citrate to avigorously stirring solutions of 0.01% HAuCl₄. After 1 min., 1 ml of asolution of composition 0.075% NaBH₄ and 1% Na₃ citrate was added.Reaction continued for five minutes. The solution of nuclei was storedat 4° C. until needed.

The first seeded colloid was prepared by refluxing 1 ml of 1%HAuCL₄.3H₂O with 100 ml of 18MΩwater with vigorous stirring. 0.4 ml of1% Na₃Citrate and 30 μl of the above described nuclei was added rapidlyand boiled for an additional 15 minutes followed by cooling to RT. Theresulting colloid was stored in a dark bottle.

A second seeded colloid was prepared by an identical method using 15 μlof nuclei instead of 30 μl.

Colloid Major Minor Std. Dev. # of Part. Nuclei 2.64 2.03 1.04 131Seeded #1 52.7 43.7 5.24 70 Seeded #2 93.4 68.0 20.0 16

FIGS. 11-13 are TEM images of the particles. FIGS. 14 is the solutionoptical spectrum of the seed nuclei. FIGS. 15-16 are the uv-vis spectraof the larger particles derivatized on glass substrates and of thelarger particles in solution.

EXAMPLE 2 Preparation of 2-layer Colloid Surfaces

Once a colloid monolayer is formed, it is possible to produce amulti-layered material by introducing a chemical linking agent and asecond layer of particle. Possible linkers include4,4′-bis-pyridylethylene, 4,4′bipyridyl, p-xylenedithiol, andmecaptoethylamine. Experimental protocol for preparation of a typicalsurface follows.

Glass slide surfaces, approximately 2 cm², were washed first in asolution of HCl:HNO₃ (3:1) followed by an H₂O rinse and cleaning in amixture of 30% H₂O₂:H₂50₄ (1:4). The cleaned slides were placed in about3 mL of a 1:10 solution of aminopropyltrimethylsiloxane (APTMS) inmethanol for 1 hour. The slides were rinsed with water and placed inabout 3 mL of a 7.0 10⁻¹² M colloid solution overnight. These particleshad the following shape: 65.8 nm major axis, and 49.3 nm minor axis. Theslides were again rinsed with water, and stored in water. A TEM image ofthe particles is shown in FIG. 17. The resulting optical spectrum isshown in FIG. 18. This surface was immersed for 5 minutes in 1 mMmercaptoethylamine in water and rinsed. This surface was then immersedin a 17 nM solution of 12 nm colloidal Au particles for 15 minutes. FIG.19 is the resulting optical spectrum.

EXAMPLE 3 Protein Coated Au Colloid Monolayers

Quartz surfaces which had been cleaned in a mixture of H₂SO₄:H₂O₂ (4:1)were derivatized in neat silane solution for 2 days. After rinsing thesesurfaces in spectrophotometric grade CH₃OH, they were placed in asolution of 3-(N-maleimidopropionyl)-biocytin (0.31 mg/ml in 0.05M Trisbuffer containing 0.1 % BSA) for 18 h. Slides were rinsed in H₂O andplaced in solutions of the Au probe described below. The protein-Auprobes had been aged for 1 hour prior to coating the slides; slidesremained in solution overnight and were removed when the colloidalparticles sedimented. Little to no coating was achieved in the first 5 hof exposure to the Au probes; optical spectra taken after the colloidsedimented indicated Au coating on all of the silane/biocytin surfacesemployed, with λ_(max)=550 nm.

Protein-coated Au probes were prepared as follows: To 25 ml of acitrate-prepared Au colloid (12 nm diameter, 17 nM) was added 0.725 mlof streptavidin (0.34 mg/ml in triply distilled H₂O) and 0.241 ml of BSA(7.24 mg/ml in triply distilled H₂O). Within 4 hours, this solutionshowed some aggregation/sedimentation of particles, but particles wereeasily resuspended with shaking the solution. However, within 24 h, theparticles had sedimented completely and could not be resuspended.

Quartz surfaces which had been cleaned in a mixture of H₂SO₄:H₂O₂ (4:1)were derivatized in neat silane solution for 2 days. After rinsing thesesurfaces in spectrophotometric grade CH₃OH, they were placed in asolution of 3-(N-maleimidopropionyl)-biocytin (0.31 mg/ml in 0.05M Trisbuffer containing 0.1% BSA) for 18 h. Exposure of this surface to astreptavidin-coated Au colloid resulted in surface formation.

FIG. 20 shows the strategies available for non-covalent attachment ofcolloids to surfaces, and how these strategies can be utilized to bindbiomolecules to metals in a biocompatible fashion. FIG. 21 compares thecoverages by covalent and non-covalent methods.

EXAMPLE 4 Detailed Experimental Protocols for the Production ofAg-coated Au Colloid Monolayers by Chemical or Electrochemial ReductionMaterials

The following materials were obtained form Aldrich: HAuCl₄.3H₂O, Na₂SO₄,trisodium citrate dihydrate, pyridine, and N,N-dimethyl4-nitrosoaniline(p-NDMA). The following organosilanes were obtained from Hüls America,Inc.: 2-(diphenylphosphino)-ethyltriethoxysilane (DPPETES),3-aminopropyltrimethoxysilane (APTMS),mercaptopropylmethyldimethoxysilane (MPMDMS) and2-(trimethoxysilyl)ethyl-2-pyridine (PETMS). KC1, HCl, and HNO₃ wereobtained from J. T. Baker. Ru(NH₃)₆Cl₃ was obtained from JohnsonMatthey. Li Silver was obtained from Nanoprobes, Inc. Methanol(spectrophotometric grade) was obtained from EM Sciences. All chemicalswere used as received. All H₂O was 18 MΩ, distilled through a BarnsteadNanopure water purification system. Single component Ag epoxy wasobtained from Epoxy Technologies, Inc and white expoxy was epoxi-patchfrom The Dexter Corporation. Glass microscope slides were obtained fromFisher Scientific, quartz slides from Technical Glass Products, andIn-doped SnO₂ from PPG Industries, Inc.

Colloid Preparation

Colloidal Au particles of 18 nm±1 nm and 12 nm were prepared asdescribed previously. The sols were characterized by opticalspectroscopy.

Surface Derivitization

Glass and quartz microscope slides were cut to 25 mm×9 mm prior tocleaning in a bath of 4 parts H₂SO₄ to 1 part 30% H₂O₂ at 60° C. Afterrinsing in H₂O₁ slides were cleaned in a bath of 3:1 HCl:HNO₃. Slideswere rinsed in H₂O and stored in CH₃OH until needed. Clean surfaces wereplaced in 2% (by volume) 2(diphenylphosphino)ethyltriethoxysilane or2-pyridylethyltriethoxysilane in CH₃OH. Solutions were shaken vigorouslyand reacted for 24 hours. Substrates were then removed from solution andrinsed thoroughly in CH₃OH, then rinsed and stored in water. Silanizedsurfaces were immersed colloidal Au for 24 hours. Surfaces were thenremoved, rinsed several times in H₂O and stored in H₂O. In-doped SnO₂was cut to an approximate area of 3.0 cm². Copper wire was adhered tothe SnO₂ with Ag epoxy. The copper wire was encased in glass tubing andsealed with white Epoxy-Patch The contact area was covered with a whiteepoxy. The electrodes were sonicated in neat reagent grade acetone, soapsolution, and H₂O for approximately 15 minutes each, then were soaked inapproximately 3 M NaOH for over 2 hours. Clean electrodes were placed inaqueous APTMS (1% by wt.) for 5 minutes. Silanized electrodes were thenrinsed with H₂O and dried for 48 hours. The electrodes were placed inthe Au colloid solution for 2 hours. Colloid-coated electrodes were thenrinsed and stored in H₂O. The other SnO₂ samples were placed in neatMPMDMS for approximately 5 hours. The substrate was rinsed with CH₃OHand air dried. The polymer-coated substrate was then placed into freshAu colloid for at least 8 hours for high coverage SnO_(2.) The Aucolloid-coated substrate was removed, rinsed with triply-distilled H₂O,and air dried.

Electrode Characterization

A 5 mM Ru(NH₂)₆Cl₃ in 0.1 M Na₂SO₄ was degassed with N₂ for over 15minutes prior to analysis. An SCE was used as a reference electrode andPt gauze was used as the counter electrode in all measurements.

Sample Preparation Li Silver Reduction

Ag was reduced onto the colloid-coated slides by immersing thesubstrates in a solution of equal volumes of LI Silver enhancer andinitiator solutions. Reaction time was varied between 5 and 30 minutes.Surfaces were then rinsed and stored in H₂O.

Ag Deposition

An electrolyte solution of 1 mM Ag₂SO₄ and 0.1 M Na₂SO₄ was degassedwith N₂ for at least 15 minutes prior to the electrochemical deposition.A Pt gauze electrode was used as a counter electrode. A cyclicvoltammogram was first done with the electrode scanning from 1.0 V to0.0 V vs SCE at 50 mV/sec. A constant potential was applied to theelectrode until a predetermined number of Coulombs had been counted. Thecyclic voltammogram and the reduction of Ag were both done in the samesolution under a blanket of N₂. The electrodes were then removed, UV-Visspectra were collected, and the electrodes were stored in H₂O.

Surface Characterization

SERS of 0.5 M pyridine/0.1 M KCI solution on the electrodes wereperformed. The potentials were obtained with a Pt counter electrode anda SCE reference electrode and were varied from 0.0 V to −0.9 V. SERS ofthe Li Silver was done on 0.5 mM p-NDMA in CH₃OH. Experiments wereperformed with 632.8 nm excitation, 7.5 cm⁻¹ bandpass, 0.5 cm⁻¹ stepsand 1 second integration.

Instrumentation

SERS spectra were obtained with a Spectra-Physics Model 127 HeNe ionlaser operated at 632.8 nm. Raman spectra were acquired with a SpexModel 1403 scanning double monochromatic with a pair of 1800 groove/mmgratings and thermoelectrically-cooled Hamamatsu photomultiplier tubehoused in a Products for Research casing. Monochromator entrance andexit slits were set for a spectral bandpass of 7.5 cm⁻¹. Spectralacquisition and grating movement were controlled via the DM3000 softwareprovided by Spex. Plasma lines were filtered out of the incident beamthrough a band pass filter (Ealing ElectroOptics.) Incident light wasfocused at an angle ˜<30° to the surface normal. Scattered radiation wascollected and collimated with a Minolta 50 mm camera lens (f/#1.2) andfocused through a polarization scrambler (Spex) onto the entrance slitsof the monochromator. Substrates were supported by Teflon block (⅓ ofsample height) in a quartz cuvette. The cuvette holder rested on ahome-built stage adjustable in all three dimensions.

Optical spectra were obtained using a Hewlett-Packard 8452A diode arrayspectrophotometer with a 2 nm spectral resolution and 1 s integrationtime. Samples were supported in the cuvettes by use of the teflon blockdescribed above.

The cyclic voltammograms of the Ru(NH₃)₆Cl₃ were done on a CypressSystems CS87 potentiostat. The cyclic voltammograms and the Agdeposition were carried out using an EG&G PAR 173/175 combinationprogrammer/potentiostat, and recorded by a NGI Servogor 790 XY recorder.Ag deposition was monitored using a Linseis L6512B strip chart recorder.The potential during the Raman scans was held constant by a BAS CV27potentiostat.

Atomic Force Microscopy (AFM) was performed on a Digital InstrumentsNanoscope III Multimode AFM in tapping mode. The samples were cut toapproximately 1 cm×1 cm and allowed to air dry. The scan size was 1.00μm, and scan rate was 0.9988 Hz. X and Y axes are 1 μm with 0.2 μmdivisions, and all the z-axis are 300 nm.

FIG. 22 show the changes in optical spectra upon electrochemicalreduction of Ag ions onto Au colloid monolayers. FIG. 23 showstheoretical modeling of the expected changes, showing that the Ag isselectively deposited on the Ag surface. FIG. 24 shows AFMcharacterization of these surfaces. FIG. 25 shows optical spectra as afunction of electrochemical potential (E) for a surface with high Aucoverage (from 0 to −1 V vs. SCE), showing that these surfaces arestable under electrochemical conditions. FIG. 26 shows the changes inoptical properties accompanying chemical deposition of Ag onto preformedAu colloid monolayers.

EXAMPLE 5

Large colloids can be stabilized by addition of polyelthyleneglycol(PEG). This does not preclude derivatization on surfaces. For example,colloidal solutions were stabilized by the addition of fresh 1% PEG (mw20,000 Fluka) until the final PEG concentration was 0.1%. The colloidalsolution was then centrifuged in glass test tubes at 3000×G for 15Minutes to pellet the colloid. The supernatant was removed and thecolloid was resuspended in distilled H₂O. This resuspended colloid wasthen adsorbed on to silanized surfaces as described elsewhere. Theresulting surface was characterized by uv-vis in FIG. 27.

EXAMPLE 6 Preparation of 2.5 nm Au Particles on thin Layers of InorganicOxides Coated with Organic Polymers.

Indium-doped SnO₂, cut to dimensions of 1 cm by 1 cm, was cleaned asfollows: surfaces were sonicated for 15 min each in soapy water, 3changes of triply distilled H₂O, and acetone. After the surfaces wererinsed in H₂O, they were soaked in 3 M NaOH for 1.5 h. Substrates werestored in H₂O until needed for further derivatization. The SnO₂ wasderivatized with alternating layers of K₂Nb₆O₁₇ and poly(allylamine)hydrochloride (PAH) as decribed elsewhere [Keller et al, JACS, 116,8817]. PAH-coated surfaces were immersed in a solution of colloidal Au(x nm±y nm) for 28 h. Substrates were then rinsed and stored in H₂O. Theoptical spectrum in FIG. 28 shows the presence of Au particles on thesurface.

EXAMPLE 7 Preparation of a 12 nm Au Colloid Monolayer on SnO₂ ElectrodePreparation

Pyrolytically deposited Sb doped SnO₂ (R=100 ohm cm²) on glass 3 mmthick was purchased from Delta Technology Lmtd. The SnO₂ on glass wascut into pieces of the size 2 cm², piranah washed and sonicated intriply distilled H₂O for 15 min. Contact to the SnO₂ was made with Agepoxy. Cu wire encased in 3 mm glass tubing was used as the lead. The Agepoxy was covered with white epoxi patch (Dexter Corporation).Electrodes were sonocated in triply distilled H₂O for 15 min. prior touse.

The clean electrodes were placed in 3 M NaOH for five hours, rinsed inH₂O, placed in 1% (by weight) aqueous APMDES for 10-15 minutes, and thenrinsed with H₂O. The electrodes were allowed to dry at room temperaturefor 2 days. The electrodes were placed in a solution of Au colloid for 3hours, 20 min., rinsed with H₂O, and stored in H₂O. An optical spectrumis shown in FIG. 29.

The Au and Ag colloid-based surfaces have many of the best attributes ofpreviously described SERS substrates (uniform particle size,electrochemical addressability, and large enhancement factors), and acombination of other features few surfaces can match (ease ofcharacterization, no geometric constraints, low cost, and very highintra- and inter-sample reproducibility). When the substrate isoptically transparent in the visible, uv-vis can be used to assessparticle coverage and spacing. These substrates are strongly active forSERS using 647.1 nm excitation, as are those prepared on glass andquartz slides. These advantages, coupled with the ability to varyparticle size, spacing, and the nature of the monolayer support suggesta rich use for these materials in fundamental and applied studies. Forexample, no theoretical model correlating SERS activity with particlesize or spacing has ever been quantitatively tested; metal colloidmonolayers should allow such experiments to be carried out. Theflexibility and control available through this method are noteworthy.Particle size, particle-polymer interactions, and the physical andchemical properties of the underlying substrate can all be manipulatedto control the nanoscale architecture produced. The solution-basedassembly protocol makes substrate fabrication routine; it also removesvirtually all constraints on substrate size/shape. Therefore, Au colloidmonolayers can be prepared on substrates that allow facilecharacterization, including macroscopic glass/quartz slides and TEMgrids. Most importantly, SERS can now be considered for applications inwhich preparation of multiple high-quality substrates was previouslyviewed as an insurmountable barrier.

Additional Applications of the Inventive Metal Colloid Monolayers AMethod for the Detection and Analysis of a Chemical Compound Dispersedin a Carrier

The improved SERS substrates, as described above, enable Ramanspectroscopy to be used as a practical method for detecting andanalysing carrier-borne chemical compounds, specifically water-bornepesticides, herbicides and other agrochemicals. In addition, they can beused to detect disinfection-generated pollutants in water, such ashaloacetic acids. They can also be used to detect, in air, anyvapor-phase compounds possessing Raman-active modes, such as explosives,fragrances and the like, in extremely low concentrations.

Background

The widespread use of pesticides in modern agriculture leads inexorablyto their appearance in water, and such contamination continues to be ansignificant threat to water quality. Accordingly, determination ofpesticide levels in aqueous samples is a necessary and ubiquitousanalytical procedure, and there is an increasing need for rapid, simple,reliable, and pollution-free methods of determining these pesticidelevels. Organophosphorous pesticides (OPPs), while not as persistent asother organic compounds used in agriculture, are nevertheless monitoredquite carefully, in large part due to their extreme toxicity. Indeed,the National Pesticide Survey (NPS) included 43 different OPPs or OPPdegradation products, and left out numerous more because of theirinstability.

A tremendous amount of published work is concerned with analysis ofthese compounds, using a compendium of analytical methods. One methodpreviously used relied on the determination of nitrogen/phosphorous(N/P)-containing pesticides in ground water by GC with an N/P detector,using methyl t-butyl ether as the final extraction solvent. For obviousreasons, there is tremendous pressure to reduce or completely eliminatethe use of organic solvents in the liquid-liquid extraction (LLE) step.Recently discussed methods for doing so include thin-layerchromatography, gas chromatography (with nitrogen-phosphorous, flamephotometric, or mass spectrometric detection), liquid chromatography(with electrochemical, uv diode array, or fluorescence detection), andinhibition of cholinesterase biosensors. In most cases, there is anemphasis on solid-phase extraction (SPE) methods using disks orcartridges, simultaneously eliminating the need for LLE and providing asample concentration mechanism; in recent reports, the SPE step has beenfused to the separation step to yield “on-line” methods. It should benoted, however, that humic acids, clays, and pH have been shown toimpact SPE extraction efficiency for several organopesticides. Moreover,several OPPs have been shown to completely degrade on SPE precolumns at4° C. A reasonable alternative to SPE is accelerated solvent extraction,a technique that reduces the volumes of conventional liquid solvents bythe use of increased temperature and pressure.

A second thrust, relevant to the entire field of pesticide analysis, isthe recent proliferation of portable instrumentation. The rationale fordevelopment of these instruments is the ability to make analyticalmeasurements on-site, a critical component of the emerging nationalwater quality monitoring program. These new instrument and protocolsplace a premium on “operational simplicity”, with the basis of theanalytical method typically resting on a single measured parameter, andsometimes even a yes-no response.

A good example of this approach is a recently introduced portableanalysis kit for a number of pesticides, including chlorpyrifos, basedon a competitive immunoassay, a technique in which labeled and unlabeledanalytes compete for binding sites on immobilized antibodies-which inthis particular technique are conjugated to magnetic particles.Separation of the magnetic particles, followed by a spectrophotometricassay of an enzyme linked to the labeled analyte [enzyme-linkedimmunoassay (ELISA)], gives an absorbance proportional to the amount oflabel. Using a calibration curve, this can be correlated to theconcentration of unlabeled analyte in the sample. A similar immunoassayfor parathion has also been reported that does not require its removalfrom hexane.

There are a number of serious drawbacks to these approaches: (i) it iswell-recognized that competitive immunoassays are not trivial to carryout, and unlike sandwich immunoassays, in which the signal isproportional to analyte concentration, competitive immunoassays yieldsignals inversely proportional to analyte concentration; and (ii)different immunoassays kits are necessary for each analyte to bedetected; no more than two uv-vis-based ELISAs can be carried out in thesame test tube, and multianalyte detection by any method is difficult.Moreover, enzyme labeled analogs and antibodies exist for only a smallfraction of OPPs in use. In addition, (iii) the extreme similarity inOPP structure (see Table II) suggests that very high specificity inmonoclonal antibodies is not possible, i.e. one OPP could act as aninterferant in an assay for another OPP, and (iv) other interferences,which are often present in real-world samples, cannot be identified insingle parameter measurements. Finally, (v) even if none of theseproblems existed, the shelf life and stability of protein-basedanalytical reagents are generally quite short. The net effect is thatwhile development of immunological approaches to on-site measurements ofpesticides is being widely pursued, the measurements must necessarily becarried by people with technical expertise; the possibility of usingthis method to determine water quality on individual farms by individualfarmers, a clear goal of the national water quality monitoring program,is slight.

It would therefore be advantageous to develop a direct method forquantitation of OPPs and other water-borne pestisides or agrochemicalsbased on surface enhanced Raman scattering (SERS), as thisultrasensitive spectroscopic technique is optimally suited for analysisof dilute aqueous solutions of organic compounds. Like its vibrationalcounterpart, infrared spectroscopy, Raman spectroscopy yieldsmolecule-specific information relating to the identity of atoms, bondstrengths, and bond angles. In other words, a Raman spectrum provides acharacteristic fingerprint for a given molecule. Few other spectroscopictechniques can match the high information content of vibrationalspectroscopy. This is to be contrasted with methods based onfluorescence and absorbance—immunoassays, for example—in which a singledatum is often used to ascertain analyte concentration. Ramanspectroscopy has a great advantage over other spectroscopic techniquesfor analysis of aqueous solutions; H₂O is a very poor Raman scatterer,in contrast to the nearly insurmountable interference H₂O generates ininfrared spectroscopy. Raman spectroscopy has not been previously usedin the analysis of water-borne pesticides, however, because Ramanscattering is inherently a very weak phenomenon, with an intrinsicsensitivity 3-4 orders of magnitude too low to detect OPPs in thesubmicromolar concentration range. The improved results achieved withthe SERS substrates formed in accordance with the present inventionenable Raman spectroscopy to be used as a practical method for detectingand analysing water-borne pesticides.

The attraction of Raman spectroscopy as a technique for pesticidedetection, and OPP detection in particular, is clearly elaborated inTable I, which lists the strong or medium Raman vibrations between 400cm⁻¹ and 1700 cm⁻¹ for 8 OPPs (whose structures are illustrated in TableII). Each of the eight species has a unique spectrum, and each has atleast one strong vibration that is spectrally well-resolved from theothers. Thus, each compound in this table can be identified directly,without any separation required. In contrast to TLC, GC, and HPLC, thisis a direct measurement in the truest sense of the word, since allcompounds are detected simultaneously on the basis of their molecularstructure. Among the many OPPs available for study, this particular setof compounds was focused on only because SERS spectra already exist: Itis stressed that this same technology is applicable to any finite set ofOPPs and to other pesticides and other classes of compounds, such ashaloacetic acids, the toxic byproducts of chemical disinfection. TableIII shows the unique Raman bands for haloacetic acids and chlorinatedphenols.

TABLE I Raman Peak Locations for 8 Organophosphorous Pesticides

TABLE II Structures for 8 Organophosphorous Pesticides

TABLE III Raman Spectra of 9 Haloacetic Acids 24- 246- TBAA DBAA BAATCAA DCAA CAA 2-CP DCP TCP Source A B C C C C B B D Band = 1 193 vs 2201 s 3 217 vs 4 233 vs 234 vs 230 vs 5 657 271 vs 6 280 vs 7 343 vs 8372 vs 370 vs 9 378 vs 379 vs 10 388 vs 386 s 11 403 s 12 412 vs 13 418vs 14 437 vs 15 449 vs 16 489 s 17 563 vs 18 614vs 19 656 vs 20 667 vs21 677 vs 22 680 vs 684 vs 681 vs 23 727 vs 729 vs 24 775 vs 25 797 vs793 vs 26 821 vs 27 835 vs 28 856 vs 29 867 vs 30 906 s 903 s 31 927 vs32 1031 vs 33 1392 s 34 1405 s Abbreviations: TBAA = tribromoaceticacid: DBAA = dibromoacetic acid: BAA = bromoacetic acid: TCAA =trichloroacetic acid: DCAA = dichloroacetic acid: CAA = chloroaceticacid: 2-CP = 2-chlorophenol: 2,4-DCP = 2,4-dichlorophenol: 2,4,6-TCP =2,4,6-trichlorophenol: vs = very strong; s = strong. Raman data forbromochloroacetic acid are not available. Darkened boxes refer tovibrations that are spectrally distinct, and whose presence can be usedto measure the concentration of a particular HAA in a mixture. Notethat: (a) for some compounds, there are multiple unique bands, each ofwhich could be used as a diagnostic; and (b) there are other, muchweaker bands in each spectrum.

It is important to note that a double monochromator and photomultipliertube detector was used to aquire the data in Table I (in 1987) and TableIII (in 1989). The instrumentation available today not only providesgreater throughput (with a single monochromator/holographic notch filtercombination) but also improved signal/noise (with CCD detectors). Thus,the detection limits for the work proposed herein will be superior tothe nanogram levels previously reported.

SERS-based detection has additional advantages relative to HPLC and GC.both of which have significant setup time, involve machinery that isdifficult to make portable, and typically require operator training. Inother words, these “direct” methods are not necessarily “simple”. Withpre-formed substrates, SERS data are extremely easy to acquire,comparable to measuring absorbance.

Another significant advantage to Raman spectroscopy over GC and HPLC isthe information content per molecule. GC and HPLC techniques identifycompounds based on their viscoelastic properties and mobilities, andyield little information: a retention time, or in the case of diodearray uv-vis detection for HPLC, an optical spectrum. Unfortunately,these properties cannot always be directly correlated to molecularstructure, particularly for compounds with such similar atomicconnectivity. Raman spectroscopy offers the advantage of multi-peakidentification, with the signal for each OPP having numerous components.In complex matrices, where there is an increased likelihood ofinterferences and/or impurities, the ability to directly identify acompound using several distinct peaks (Table I) is a huge advantage.This property is especially important for compounds like OPPs thatdegrade rapidly; characteristic vibrations can be used to identifydegradation products, without knowing the degradation chemistry.Similarly, characteristic frequencies can be used to identify otherclasses of pesticides. In contrast, an unknown compound cannot beidentified by GC or HPLC solely on the basis of its retention time. Itshould be noted that phenylamide pesticides have been quantitated byResonance Raman scattering in solution, but this requires derivatizationwith an intensely absorbing azo dye derivatives. Such an approach wouldnot be practical for OPPs, which lack the appropriate chemicalfunctionality.

A final advantage to SERS is that it is an extremely general method; ifanalyte adsorption can be induced, it can be used for any water analysiswithout any changes in equipment or configuration. As water qualitytakes on increased significance in society, and as the number ofdifferent compounds in water that must be monitored increases, this isan important consideration.

SERS behavior of colloid monolayers can further be controlled byadjusting the intrinsic properties thereof. It is known, for example,that the dielectric constant of Ag make it more enhancing than Au forvisible light excitation. However, over the 10-100 nm scale, Au colloidsare more monodisperse than Ag. Thus, in a preferred embodiment of theinvention, the strengths of each metal are exploited by usingautometallography, a technique whereby Ag⁺ is selectively reduced ontopre-existing Au particle surfaces. Thus, submonolayers of monodispersecolloidal Au, sufficiently separated so as to be only weakly SERSenhancing, are overcoated with Ag. The resulting particles are largerand therefore exhibit reduced interparticle separation. Particle arraysformed thereof possess Ag-like optical properties and enhancementfactors. Testing with Ag-coated Au confirms that the SERS intensityachieved with such particles are within a factor of two of the signalobtained with pure Ag colloid, and within a factor of ten of the bestroughened Ag electrodes. These results indicate that these surfacespossess exceptional enhancement characteristics.

While SERS is capable of providing an enormous stream of data, it willbe necessary to rapidly convert the data into a usable form. One way todo so is to use artificial neural networks (neural nets) for this task.A growing body of work describes the use of neural nets for spectralrecognition, quantitation, or classification. Applications to Mossbauer,laser-induced fluorescence, and mass spectra are notable. A particularlyintriguing implementation appears in commercial instrumentation for odordetection.

Neural nets have previously been used in several capacities with regardto pesticide physical properties. These include an estimation of soilpartition coefficients based on chemical structure, a model forhalf-lives, and an assessment of toxicity. More importantly, experimentshave been carried out to classify pesticide IR spectra intoorganophosphorous and non-organophosphorous classes with neuralnetworks. The same group also used a three-layered neural network forautomated interpretation of pesticide IR spectra. These results, coupledwith demonstrated recent successes of neural networks for multiparameterspectral analysis, indicate that the proposed neural net-basedidentification and quantitation of OPPs is possible.

Establishment of a non-separation based analytical method on samplescontaining mixtures requires three stages of experimentation. In thefirst, detection limits are established for each analyte individually.In the second, detection limits are established for mixtures. Finally,signal calibration is required for mixtures with differingconcentrations.

Measurement and optimization of individual detection limits is astraightforward task. The detection limit is defined according toconvention as the amount of sample producing a signal/noise ratio of 2.A tremendous advantage of Raman over separation-based methods is theremoval of flow-induced limits on integration times. In other words,since the analyte is not moving, one can integrate the Raman spectrumfor arbitrarily long times. For CCD detectors, background noise isvirtually nonexistent, but practical limitations on integration timesare about 15 minutes. Nevertheless, this number should be compared tojust the few seconds of integration time available for a chromatographicdetector. There are two fundamentally different approaches to measuringSERS spectra of OPPs, each with different detection limits. In Method A,the solution is made basic to insure complete deprotonation of OPPs. Asmall aliquot (50 μL) is applied to an Ag/Au colloid monolayer on asubstrate of 1.0×1.0 cm dimensions. The hydrophilic nature of thesesurfaces allows an acceptable degree of wetting; the resulting highsurface area/volume ratio leads to rapid evaporation of excess water.Alternatively, the substrate can be mildly heated to promoteevaporation. The sample is then placed in front of the laser beam andthe SERS spectrum is measured. In Method B, a glass slide containing acolloid monolayer is immersed in the treated H₂O, and the SERS spectrumis acquired in situ. Below are estimates for the detection limits foreach method.

Method A

Assumptions

(1) Detection limit=10⁻⁵ monolayers of adsorbate

(2) Sampling area=focused laser spot size ≈2.0 mm×2.0 mm

(3) Molecular area ≈40 Å²/molecule

These assumptions are completely reasonable. (1) Single monolayers ofadsorbates can be detected by Raman spectroscopy using CCD detectors onunenhanced surfaces. With an enhancement factor of 10⁶, it thereforefollows that 10⁻⁶ monolayers can be detected on SERS-active surfacesusing a CCD. Thus, a detection limit of 10⁻⁵ monolayers is conservative.(2) Focusing a laser through a microscope objective allows Raman spectrato be obtained from spot sizes approaching 1 μm in diameter—thus the 4mm² sampling area is simple to achieve. (3) The value of 40 Å²/moleculetranslates to 2.5×10¹⁴ molecules/cm², a commonly cited value formonolayer coverage of small adsorbates on surfaces.

Sample Calculation

 Let MNDAM=the minimum number of detectable adsorbate molecules=0.00001monolayers×0.04 cm² sampled×1 molecule/40 Å²×10¹⁶ Å²/cm²=1×10⁸ molecules

Only the actual volume above the sampled area is relevant to thedetection limit, and this is the sampled area/total area multiplied bythe total volume, i.e. (0.04 cm²/1 cm²)×50 μL=2 μL.

The presence of 10⁸ molecules in a volume of 2 μL corresponds to aconcentration of 83 pM. For a species with MW=200 (around the average MWfor the 8 OPPs under consideration), this is equivalent to 16.6 ng/L.Assuming all 8 OPPs are present in equal concentration, the detectionlimit will be increased by a factor of 9. Thus, a simultaneous detectionlimit of 149 ng/L=0.15 μg/L is anticipated. This number is equivalent todetection limits reported for GC, but without any samplepre-concentration. In other words, direct adsorption to SERS substrateseliminates the need for sample concentration using SPE or liquid-liquidextraction. This is a significant advantage to this method.

Method B

Assumptions

Whereas Method A has the advantage of effectively adsorbing everymolecule in a given volume of solution through increased evaporation, inMethod B one is constrained by K, the equilibrium constant foradsorption. This parameter defines the surface coverage for a givensolution concentration (c). In addition to the assumptions delineatedabove, the calculation for in situ detection limit will assumeLangmuirian adsorption (i.e. no interaction between particles). Thus,q/(1−q)=Kc, where q is fractional surface coverage. In the absence ofconcrete data regarding attraction or repulsion between particles, it isa reasonable approximation to assume a Langmuirian adsorption isotherm.A value of K=10⁵ on Ag is likely an underestimate for two reasons:first, OPPs are sparingly soluble, which dramatically increases theiraffinity for surfaces; and second, most of the compounds in Table IIcontain sulfer, which binds very strongly to Ag and Au.

Sample Calculation

Using q=10⁻⁵, c=q/[K(1−q)]=10⁻⁵/[10⁵(0.99999)]=1×10⁻¹⁰ M. With MW=200,and all 8 OPPs present, this corresponds to 0.18 μg/l, a value similarto that for method A, abut again without the need for pre-concentration.

It is important to note that Method B essentially corresponds to an insitu method, and as such, has important ramifications. A protocol thatconsists of withdrawing 2 mls of sample, placing it into a uv-viscuvette, adding a colloid-coated glass slide, and pressing a button on acomputer console is exceptionally attractive. If an internal standard isdesired (vide infra), this could be added to the cuvette. Furthermore,if the colloid monolayer technology could be transferred to an opticalfiber (a likely scenerio, given the success with oxidic surfaces such asglass, quartz, SnO₂, and Al₂O₃), true in situ monitoring can beachieved. Raman spectroscopy using optical fibers is well-established,as is its use to monitor industrial processes; it would be invaluable tomonitor speciation of pesticides in real time.

Inherent to the inventive methods are four possible problems:non-systematic intensity variations, the possibility of interferences,the need for adsorption of analyte, and surface saturation-inducednonlinearities in signal. However, each of these can bestraightforwardly addressed.

Addition of an internal standard eliminates any concerns regardingsurface and/or intensity variation. Any one of hundred of compoundspossessing a Raman spectrum distinct from those of the OPPs wouldsuffice. So that adsorption properties would be similar, structuralsimilarity is desirable; thus, fluorinated phenols are a logical choice.Small volumes of known concentrations would be added prior tomeasurement.

The most likely interferences in SERS are adsorption of inorganic anions(HO³¹, HCOO³¹, etc.) Their influence can be minimized by acidifcation;at pH 5, [HO⁻]=10⁻⁹. Thus a simple route to identify (and eliminate)undesired adsorbates is to acquire Raman spectra at several pH's. Halideanions are bound more tightly, but each have only one band in the Ramanspectrum, so that their net effect is essentially limited to a reductionof available OPP binding sites. If this leads to significant problems,it can be overcome by introducing an element of molecular selectivity atthe Au/Ag interface. The introduction of organothiol self-assembledmonolayers (SAMs) onto SERS-active surfaces increases selectivity forthe desired analytes, and that partition from solution into the SAM filmcan be quantitated. A similar strategy can be adopted with regard to theinventive method using HSCH₂CH₂OH or HSCH₂CH₂NH₃ ⁺. Monolayers of thefirst molecule should preferentially hydrogen bond with the P=S or P=Ofunctional group relative to H₂O, HO⁻, or halide. Note that directanalyte adsorption is not necessary for SERS: enhancement by theso-called electromagnetic effect—which accounts for 10⁴ of the 10⁶overall enhancement—is still quite substantial 10 Å from the surface.

A legitimate concern is the presence of organic carboxylate impurities.These can be avoided in the preparation of colloid monolayers, but maybe present in treated waters (a disadvantage to non-separation basedmethods). Fortunately, if such compounds are present, they can be easilyidentified by their Raman spectra. Considering the purity of the watersunder consideration, it is anticipated that this will not be a seriousproblem.

All 8 OPPs will strongly adsorb to Ag, but do not adsorb to equalextents. If two species are present in equal concentration in solution,but one adsorbs with greater affinity, the true concentration of thepoorly adsorbed species can easily be masked. Fortunately, thestructures of the OPPs fall into two narrowly-defined categories(phosphotriesters and phosphothiotriesters). Within each structuralclass, the equilibrium constants will vary little. Any non-zerodifferences between the equilibrium constants for adsorption can bemeasured and factored in. This process is further described below in thediscussion of mixture analysis.

A final potential problem is saturation of the surface-binding sites: athigh OPP concentrations, this could lead to a non-linear response.Inaccuracies can be avoided by diluting the original OPP-containingsolution by 10-100 when peak intensities exceed 50000 counts per second.If there is no appreciable decrease in signal intensity, the surface islikely saturated. A good procedure for dealing with this is to discardthe substrate, dilute by a factor of 500-1000, and repeat themeasurement with a new substrate. In general, it is desirable to runeach sample at three slightly different concentrations undernon-saturating conditions, particularly since the measurement processitself is so simple. This will yield more statistically valid data.

To best understand the complexities of mixtures, the ranges of K foreach class of OPP should be determined. Using a microcalorimeter,adsorption isotherms can be generated, from which K can be calculated.With values for K in hand, mixtures are prepared, and the measuredintensities compared to those predicted. If they are identical, then bymeasuring all K's, it is possible to know the exact surface distributionof a prepared mixture a priori. More likely is the case where thepredicted and observed differ, in which case it will be necessary togenerate calibration curves manually. This requires substantial effort,and can constitute a critical component of the protocol.

Alternatively, the method of standard addition can be used instead ofcalibration curves. In this approach, known amounts of OPPs are added tosamples containing unknown; the x-intercept that results from the linearplot of signal vs. amount added gives the concentration of unknown.

Thus, as described above, surface enhanced Raman scattering (SERS),coupled to a neural net can be used to identify and quantify water-bornepesticides and other compounds, such as haloacetic acids, with detectionlimits below 0.2 μg/L. This can be accomplished without the need forextraction or separation. This method makes use of Au, or preferablyAg-clad Au colloid monolayers, used in combination with SERSinstrumentation. In contrast to existing methods, the method of theinvention is remarkably simple to carry out, and provides directspectral identification for each compound. The inventive technique canbe used alone, or interfaced to existing separation methods. Further,the inventive method makes on site detection of water-borne pesticidespossible.

Colloid Monolayer-Based Detection In Separation Experiments

SERS can also be used as a detection method in a separation experiment.A default protocol would thus be to do GC or HPLC with SPE, adsorbingthe eluate directly onto colloid monolayers. Using a micromanipulator toraster the surface across the solvent exit port will thus preserve thechromatogram spatially on the SERS substrate, which can then beaddressed in a site-by-site fashion by SERS. For HPLC, this is atechnically trivial task. The task is slightly more complex for GC;nevertheless, a SERS-based GC detector has been described. In any case,coupling HPLC or GC with SERS eliminates any problems mixtures engender,while considerably improving the detection limits currently availablefor each.

SERS can also be used in conjunction with capillary electrophoresis(CE). The basic idea behind SERS detection in CE is to spread the eluateonto an Au or Ag (or Ag-coated Au colloid monolayer on conducting SnO₂and then interrogate the surface using microRaman. FIG. 30 illustratesthis approach. An electric field is applied between one solventreservoir and a conductive substrate, such as a doped SnO₂ surface.Alternatively, the tip of the capillary can be metallized, and theelectric field applied thereto, in which case a non-conductive substratecan be used. One end of the capillary is placed in the solventreservoir, and the other end is contacted to the Au colloid monolayer.The substrate is then rastered by the capillary. As compounds A, B, andC elute, they are deposited at different spots on the surface.Elimination of diffusion and forced adsorption can be simultaneouslyeffected by solvent removal, and MPMDMS(mercaptopropylmethyldimethoxysilane)-based Au colloid surfaces can beheated to 70° C. with no effects on optical or SERS behavior. Onceimmobilized, the substrate is rastered across a focused laser and thespectra are recorded using a CCD. Like recently-popularized direct uvfluorescence detection schemes for CE, no pre- or post-columnderivatization is needed. However, this method is not restricted to uvfluorophores and works for all Raman-active molecules.

It has previously been demonstrated that a capillary can be rasteredwhile in operation and that Au and Ag colloid monolayers have a highdegree of reproducibility from area to area and within a given sample.Therefore, several runs can be carried out on a given substrate.Alternatively, the sample can be discarded and another used—the value ofthe Au on a microscope-slide sized substrate is a fraction of one cent.In other words, these substrates are disposable. SERS has previouslybeen demonstrated using a microRaman apparatus with attomole masssensitivity, and detector sensitivity has improved considerably sincethen.

A possible pitfall in the device shown in FIG. 30 is the presence ofnon-uniform fields on the colloid monolayer surface, resulting inanalyte migration. If this occurs, it can be circumvented in two ways.The simplest approach is to make electrical contact to both ends of theSnO₂ substrate, eliminating any potential drops across the surface.Another alternative is to metallize the outer surface of the capillaryend and make contact to it.

A conservative estimate of the detection limit can be calculated using a5 μm×5 μm spot size and a SERS signal/noise (S/N) ratio of 1000:1. Ramanmicroprobes allow focusing down to spot sizes down to 1 μm×1 ρm, andwith the CCD detector now available, a S/N of >5000:1 is attainable.With a monolayer coverage of 10¹⁴ molecules/cm², one calculates 25μm²×10¹⁴ molecules/cm²×1cm²/10⁸ μm²=25×10⁶ molecules in the laser beamfor a full monolayer. Factoring in a detection limit where S/N=2, thiscorresponds to 50,000 molecules.

Surface-Confined Protein/Au Complexes and the Use Thereof as Biosensors

Studies indicate a future trend in the medical field will lead to anincreased use of implantable “biochips”, responding to biosensoranalysis of health-related biological markers via controlled release ofsurface-confined drugs. Before such technology can be practically used,however, dramatic advances are needed in biosensor architecture: thedistinct fabrication chemistries and different readouts (photons,current, frequency, etc.) of the present devices preclude integrationinto miniature, multi-sensor arrays. What is lacking is a uniformmechanism for transducing a ligand binding event or a solutionconcentration into a measurable quantity. In view of this need, as wellas for the need to replace immunoassays with more rapid methods,Applicants have developed a series of biosensors based on complexesbetween biomolecules and colloidal Au nanoparticles. It has been foundthat appropriately sized colloidal Au nanoparticles can be used asnon-denaturing photon and electron “antennae” for biomolecules, allowingbiomolecule/Au colloid conjugates to be used as building blocks for avariety of sensing mechanisms. More specifically, this aspect of theinvention is directed to biomolecule/Au complexes that can be used todetect biological ligands (either small molecules or proteins) throughmeasuring of binding-induced changes in electrical resistance, using thepreviously described self-assembled metal colloid layers.

Discontinuous conductors surrounded by insulators are an imporant classof materials that include evaporated thin metal films andparticle-loaded polymers. At low conductor concentrations (p), thesematerials do not conduct electricity; at high p, conductivity approachesbulk values. At a certain critical concentration (p_(c), the percolationthreshold), dramatic increases in ac and dc conductivity are observed.In this threshold region, small changes in insulator volume (whichdecrease conductor concentration) lead to large changes in filmresistance. Accordingly, small molecules that induce swelling inpolymers can be detected by measuring the resistance changes inpolymer/conducting particle composites. Similarly, volume changes inpercolating Au/antibody films, induced by antigen:antibody complexformation can be detected by simple resistance measurements which canthen be converted to analyte concentration data. Related implementationsinclude detection of receptors using arrays of Au particles harboringsurface-confined ligand, and detection of oligonucleotides usingsurface-confined complementary fragments.

As was previously noted, Au colloid-based macroscopic surfaces aresimple and inexpensive to prepare, stable, amenable to assembly in largenumbers and in any geometry, extremely reproducible, capable of furthermodification (i.e. Ag deposition to yield Ag-clad Au), SERS-active, andelectrochemically addressable. Further, surface confined protein/Aucomplexes retain bioactivity, a property of the utmost importance forpotential sensor applications.

An important feature of the Au colloid-based substrates is the controlof feature size, spacing, and substrate coupling available throughself-assembly. Size control is simple, as descibed above. Sincesolution-based surface formation is based on particle diffusion,manipulation of interparticle spacing can be achieved through a detailedunderstanding of the kinetics of surface evolution. In addition,tremendous variation in coupling between colloidal submonolayers and theunderlying substrates is available through modulation of the spacer filmthickness.

These parameters are crucial in determining how metal surfaces interactwith proteins. Numerous studies using a variety of spectroscopictechniques have shown that direct protein adsorption on macroscopicmetal surfaces leads to partial or complete denaturation. The mostlikely, process for this denaturation is through formation ofthermodynamically favorable protein-surface contacts that lead toreduced conformational flexibility and, eventually, to unfolding. Thedenaturing properties of macroscopic metal surfaces pose a seriousproblem for biosensors, which typically require metals to transducebiological signals.

It has been found that, unlike macroscopic metal surfaces, smallcolloidal Au particles do not denature proteins as the high curvature ofthe nanoparticles minimizes the surface area available to the proteins.In fact, colloidal Au particles of ≦5 nm in diameter are the same sizeas the proteins themselves. In contrast, the diminished curvature oflarger particles leads to approximately the same characteristics notedwith macroscopic Au surfaces, and thus, it is generally accepted thatlarger Au particles, as well as aggregates of smaller Au particles, havean increased tendency to denature proteins.

The surface confined protein/Au complexes can be formed by exposing thesurface of a previously formed Au colloidal monolayer to protein.Alternatively, a pre-formed protein/Au complex can be immobilized fromsolution onto a substrate. The latter method is preferable as it allowsthe particle:protein ratio and the biological activity to be ascertainedprior to immobilization. The ability to prepare macroscopic, bioactivemetal interfaces of well defined structure is the key step to a uniformbiosensor architecture, and when coated with protein receptors, orantibodies, such surfaces form the basis for a novel type of biosensorbased on ligand binding-induced conductivity changes.

While ligand-induced changes in the dielectric constant of protein filmshave been proposed as a sensing mechanism, and changes in solutionconductivity are routinely used to measure cell growth, the use ofbiopolymers as molecule-sensitive insulators in composites with metalparticles has not been previously suggested. Such technology washeretofore unavailable as only Au nanoparticles combine the high degreesof conductivity and biocompatibility needed for operation of this typeof biosensor, and until the present discovery of a self-assembledcolloidal Au monolayer, there was no mechanism for the preparation ofmacroscopic, well-defined two or three dimensional particle-based filmshaving the degree of coverage required for conductivity. The twodimensional particle based films display a p≈0.16. The inventive method,however, allows the interparticle spacing to be decreased (by, forexample, coating the Au particles with Ag ) to such a degree that a p ofabout 0.60 is achieved. Alternatively, multilayers with p≈0.4 to 0.6 canbe formed. In the three dimensional regime, these are conductive.

The varying of particle spacing has been found to have a profound impacton the electronic and electrochemical properties of surface-confinedcolloidal Au films. For example, in measuring the sheet resistivity(measured by the four-point probe method) of a 12-nm diameter colloidalAu film on glass as a function of absorbance at 520 nm, in whichincreased absorbances (which correspond to decreased spacing) wereeffected by repetitive exposure of the Au colloid-derivatized surfacesto HOCH₂CH₂SH and colloidal Au in solution led to an abrupt five orderof magnitude drop in resistivity as the absorbance increased from 0.4 to0.6. Because this absorbance range corresponds to relatively closepacked monolayers (as opposed to multiple layers), the data demonstratesthat sensitive resistance measurements on monolayers of protein/Aucomplexes can be performed. Further, the insulator-conductor transitionis extremely sharp compared to those obtained with vapor-deposited thinAu films, demonstrating that protein/Au complexes assembled to thepercolation threshold are extremely sensitive to volume changes.

The colloidal Au monolayers described above have the properties of a twodimensional system, surrounded by air. Because three dimensionalcomposites of conductive particles and insulating polymers display amuch lower Pc, the use of Au colloid multilayers is prefered to amonolayer arrangement. Therefore, there is described below a method forthe stepwise assembly of conducting Au colloid multilayers fromsolution.

When a Au colloid monolayer on 3-aminopropyltrimethoxysilane(APTMS)-coated glass is exposed to a molecule having two functionalgroups that bind to or induce flocculation of colloidal Au{2-mercaptoethanol, 2-mercaptoethylamine, HS(CH₂)₆SH, APTMS,polyvinylpyrrolidone, etc.}, immersion in colloidal Au leads toadditional particle binding. Repetitive dips (5-15 minutes for thecrosslinker, 5-60 minutes for the colloid) into bifunctional organiccrosslinker and colloidal Au solutions (with copious rinsing betweenimmersions) rapidly leads to formation of Au colloid multilayers. Notethat no interlayer registry of particles is expected. This method isvery general, and can be used with colloidal Au nanoparticles of anydiameter, although the lower particle concentrations for standardpreparations of both large and small particles lead to longerderivatization times.

Tapping-mode atomic force microscopy (AFM) has been used to monitor thegrowth of colloidal Au films. Images obtained after 2 and 5 exposures to2-mercaptoethanol/colloidal Au indicate a porous geometry,with neitherclose-packed coverage within a layer (in accord with previous findings)nor registry between layers (FIG. 31). Importantly, these AFM imagesclearly indicate large areas of uninterrupted particle contact afterjust two 2-mercaptoethanol/Au treatments, suggesting the possibility ofconductive paths.

Further support for metal-like attributes comes fromvisible/near-infrared optical transmission spectra as function ofparticle coverage for samples immersed between 1-12 times in2-mercaptoethanol/12-nm Au. The spectra for low coverages match thosepreviously obtained for 2-D Au arrays or small Au aggregates insolution: strongly absorbing in the visible and transparent in thenear-infrared (FIG. 32). As the particle coverage increases,near-infrared transmission decreases markedly and becomes only weaklywavelength dependent. Similar behavior is seen in evaporated thin Aufilms, which also exhibit linear plots of transmittance vs. p at 1500 nmor 2500 nm. The intersection point of these lines (i.e. the point atwhich % T is wavelength-independent) corresponds to p_(c). In Au colloidmultilayers, a similar plot shows no such intersection (FIG. 33).

Nevertheless, at sufficiently high coverages, Au colloid multilayersundergo an insulator-conductor transition. A plot of DC resistance(plotted on a log scale) vs. particle coverage shows that samplescontaining 15×10¹¹ particles/cm² are insulating, with a resistanceapproaching 10⁸Ω. Increasing particle coverage leads to an enormousdecrease in resistance; when the particle coverage is doubled to30×10¹¹/cm², the resistance drops below 100Ω, a change of almost 6orders of magnitude (FIG. 34). Considering the extremely porous natureof these films, and the presence of organic adsorbed 2-mercaptoethanolon the particles, these films are remarkably conductive: the mostconductive sample (12 treatments) has a resistivity of 5.1×10⁻⁴ Ω-cm, avalue roughly 1/200 that of pure Au of the same geometry. Au colloidmultilayers on glass substrates are thus more conductive than In-dopedSnO₂, for example, and can be used for routine voltammetricmeasurements. Electron transfer between particles likely occurs byactivated hopping between nearest neighbors, as previously observed forgranular metal films.

Interestingly, the Au volume fraction does not change with increasingparticle coverage, evidenced by a linear plot of coverage vs. number ofimmersions in Au (FIG. 33). In other words, each step in the multilayerassembly adds the same amount of Au; the slope of the line(2.66×10¹¹/cm) corresponds to p=0.38. This value is below that neededfor 2-D percolation, but sufficient for 3-D percolation, suggesting thatthe dramatic decrease in resistance results from establishment ofmultiple 3-D percolation paths.

Devices for determining the presence and/or concentration of abiomolecule, with a biosensor comprising a receptor, antibody, orbiocomplement-coated Au colloidal layer can be constructed in the samemanner as devices using the conventional discontinuous conductor ininsulator-based sensors. As shown in FIG. 36, such a device couldinclude, for example, the inventive biosensor 101, means for applying avoltage across the biosensor 102, means for measuring an electricalcharacteristric (e.g. resistance) of the biosensor 103 and means forconverting resitivity data to analyte (antigen) concentration 104. Analternative, but equally viable implementation of the device shown inFIG. 36 involves measurement in air. The present technology is notlimited to any specific class of biomolecules and is applicable tobiosensors for detecting any biomolecule for which there is acorresponding binding partner.

Binding of a colloidal Au particle monolayer to a preformed Au colloidmonolayer or multilayer changes the effective film thickness, a factthat can be used to fabricate a biosensor based on changes in surfaceplasmon resonance (SPR) signals.

To wit, a Au colloid monolayer or mulitlayer of insufficient thicknessto achieve resonance (at a given excitation wavelength/angle ofincidence) is coated with an antibody. This surface is then exposed toanalyte (antigen). After rinsing, the sample is exposed to a Au colloid:antibody complex (the same antibody as bound to the surface). Binding ofthe Au:antibody to the surface-bound antigen leads to an increase in Aufilm thickness leading to resonance. Thus, in contrast to a conventionalSPR method, which measures small, binding event-induced changes inreflectivity of a thin Au film under resonant plasmon excitation, thebinding event described above leads to an enormous increase inreflectivity. This novel approach has several advantages. First, the“sandwich” immunoassay approach is widely used in immunochemicalanalytes, including enzyme-linked immunoassays (ELISAs). Thus, there islittle doubt that chemistry described above will obtain. Second, thisapproach can be used to sense small molecules, not routinely possiblewith current SPR technology. Third, the sensing chemistry is general forany analyte B in an A-B-C ternary complex. For example, A and C can benon-complementary strands of DNA attached to surface-bound and solutionphase DNA, and B can be an oligonucleotide (to be sensed) with ends thatare complementary to A and C.

Optimization of the Surface Characteristics of the Metal ColloidMonolayer

Preferably, for any specific application of the metal colloidmonolayers, the surface characteristics, i.e., Au particle coverage(density), Au particle size, Ag cladding thickness, will be optimized.To determine the optimal surface characteristics, a solution-basedcombinatorial strategy for synthesis of surfaces exhibitingnanometer-scale variation in mixed-metal composition and architecturehas been developed. With this method, continuous or stepped gradients inthe size and number density of surface features can be generatedsimultaneously on different regions of a single substrate. In order todemonstrate this technique, the surface enhanced Raman scattering (SERS)response of a glass slide upon which the coverage of colloidal Au andthe extent of Ag coating were varied and optimized. Over a 2 cm×2 cmsample area, changes in enhancement factor (EF) spanning four orders ofmagnitude were realized, and the changes in nanostructure responsiblefor these effects were determined using atomic force microscopy (AFM).The results show significant changes in SERS EFs over small alterationsin surface morphology, demonstrating the power of solution-basedcombinatorial approaches for optimizing the characteristics ofmixed-metal materials.

The general combinatorial assembly approach is shown in FIG. 37. Thenumber of colloidal Au particles bound to functionalized surfaces can bepredicted from immersion times in colloidal solution using initial(time)^(½) kinetics. Thus, when a 3-mercaptopropyltrimethoxysilane(MPTMS)-coated glass slide attached to a motorized translation stage isimmersed at a fixed rate into a 17 nM aqueous solution of 12-nm diametercolloidal Au, a gradient in particle coverage is generated in thedirection of immersion. After sample rotation by 90°, fixed-rateimmersion into a Ag⁺-containing solution designed to specificallydeposit Ag on Au leads to a gradient in particle size over the newimmersion direction. The resulting surface exhibits a continuousvariation in nanometer scale morphology as defined by particle coverageand particle size that results from known, repeatable immersionconditions. In preparing the gradient surface, a stepper motor with a 2°step yields a 3 μm translation. Over a 3 cm×3 cm region, it is thereforepossible to generate 10⁸ different, 9 μm² nanostructures. Macroscopicregions with uniform morphology can be prepared by very rapid successivesteps followed by a long delay. These stepwise gradients are especiallyuseful in examination of bulk changes in physical properties. Forexample, optical changes linked to growth of the characteristic Agplasmon band at 400 nm with increasing Ag coverage, evident to somedegree in continuous gradient samples, are clearly visible as squareregions with uniform shading. FIG. 38 shows optical spectra of 9different Au/Ag coatings, showing evolution of the Ag plasmon mode.

To demonstrate the SERS screening approach, the SERS response onspatially discrete areas of a 2.5×2.0 cm continuous gradient substratewere determined; the results are shown in FIG. 39. The data clearly showregions where the SERS intensity was maximized. A second example in FIG.40 shows similar data plotted in a different way. Here, increasing SERSintensity is denoted by increasing depth of shading on a grid, with themost intense spectra corresponding to the darkest squares.

An example of combinatorial signal optimization was determined asfollows. SERS signal for adsorbed p-nitrosodimethylaniline (p-NDMA) wasmeasured over a 2 cm×2 cm sample exhibiting continuous gradients in Aucoverage and Ag cladding thickness. The immersion times for samplepreparation (0-6 hr for the Au gradient, 8-22 min for the Ag gradient)were established by determination of the most enhancing region from aninitial, large gradient SERS screen on a 2 cm×2 cm sample (0-12 hr forAu, 0-30 min for Ag). FIG. 41 shows a spatial map of thebackground-corrected SERS intensity for the phenyl-nitroso stretch at1168 cm⁻¹. Intensity trends from 15 initial spots at 0.4 cm intervalswere used to locate the most SERS-active area on the sample. Furtherinterrogation at smaller intervals revealed a region that was at least10⁴-fold more enhancing than the least SERS-active domains of thesubstrate. The location of this region, a narrow diagonal band ofmoderate Au particle coverage and Ag coating, is surprising, suggestingthat maximum EFs are obtained at relatively fixed interparticle spacing.Such spacing can be achieved either by a thin coat of Ag onclosely-spaced Au particles, or with larger Ag coatings on particlesexhibiting slightly larger spacing.

AFM was used to determine the nanometer-scale morphology at positions ofinterest (FIG. 42). In FIG. 42, the image labels A-D correspond torepresentative morphologies at locations marked by shaded squares inFIG. 41. Sampling positions were chosen to highlight variation in Agcladding thickness (i.e. to span the vertical gradient) as opposed tovariation in number density of particles. Accordingly, differences inparticle coverage in images A, B and D are not significant, andrepresent positional uncertainty in the horizontal (coverage) gradient.Immersion time in Ag⁺ increases in the order D>C>B>A (A=0 min Ag), butSERS intensity follows the pattern C>B>D>A. Section analyses reveal thatmean particle size increases from A to C as expected, but decreases inD. Two related effects contribute to this result; the formation of smallAg particles between Ag clad Au colloids, and the loss of monolayerstructure (i.e. formation of multilayers) upon extended Ag exposure.Evidently, interconnection of closely-spaced particles to yield surfaceswith a morphology resembling roughened electrodes has a deleteriouseffect on SERS.

The wavelength dependence of the SERS intensity is shown in FIG. 43. Thedata show that the highest SERS enhancement factors were obtained using647-nm excitation, even though the v⁴ Raman effect favors a lowerexcitation wavelength. Using this wavelength, the nanostructure wasoptimized over two iterations, as shown in FIGS. 44A and 44B. Thesefigures show position-dependent SERS intensity maps. The results of thefirst map were used to optimize the maximum signal in the second map.FIG. 44C is a SERS intensity map generated using 28-nm diameter Aunanoparticles. FIG. 44C shows that the use of the larger core particlesleads to an increase in overall signal, as evidenced by the differentintensity scales.

A stepped gradient synthetic protocol was prepared that allowed a 10×10array of separate glass slides to be simultaneaouly derivatized andcharacterized. Using this apparatus, a 5×5 array was selected forfurther analysis. The 3D bar graph shown in FIG. 45 depicts particlecoverage and height measurements from several AFM images collected froma 5×5 array of Ag-clad-Au samples. The Au particle diameter beforeexposure to Ag was 14 nm. Immersion times were chosen using previouscombinatorial data as a guide. The Z-scale on the graph shows the numberof particles/μm², reflecting the increasing Au particle coverage withexposure time. The number written on each bar is the average particleheight on the sample, as determined by AFM section analysis. Asexpected, the particle height increases with continued Ag platingsolution exposure.

Representative AFM images, uv-vis spectra, and SERS spectra from thisdata set are shown in FIGS. 46A and 46B, FIGS. 47A and 47B, FIGS. 48A,48B and 48C and FIGS. 49A, 49B and 49C. For Ag₁, Ag₂, and Ag₃, thesamples were exposed to colloidal Au for the same time, thereforedifferences in optical spectra and SERS intensity arise from differentamounts of Ag cladding. All three optical spectra exhibit a typical Mieabsorbance at ˜430 nm for Ag, and a collective plasmon band (˜650)indicating at least partial electronic coupling between particles.Similarly, for Au₁, Au₂, and Au₃, coverage of Ag is fixed. The data showthat changes in SERS intensity from non-optimal Au colloid coverage aresignificantly more dramatic than those seen by varying the Ag cladding.

In another example, images and spectra were collected from acombinatorial sample series using 28-nm diameter colloidal Au. Eachsample was exposed to colloidal Au for the same amount of time. Totalparticle coverage needed to achieve high SERS signals is significantlylower when using larger colloid cores. Optical spectra exhibit an Au Mieresonance at ˜530 nm in addition to the collective plasmon and Ag bandsfound in the Au/Ag optical spectra having smaller Au particles, as shownby FIGS. 50A and 50B. As shown, SERS intensity is significantlyincreased with larger Au cores (FIG. 50B). AFM for these samples areshown in FIGS. 51A, 51B and 51C.

The ability to vary surface nanostructure combinatorially from solutionis extremely important for optimizing the surface characteristics ofself-assembled metal colloid monolayers for SERS, surface plasmonresonance, sum frequency generation and electrochemistry, wheresubstrate composition and topography are critical. The utility of thisapproach in materials synthesis is further augmented by the widespreadavailability of dispersible metal-containing nanoparticles and thenumerous routes to metal deposition from complexed metal cations.Moreover, recently developed Raman imaging methods are intrinsicallymore efficient than the point-by-point mapping method described herein.With tunable filter imaging spectrometers and charge-coupled devicedetectors, it should be possible to simultaneously probe up to 10⁶sample structures.

Thus is described my invention and the manner and process of making andusing it in such full, clear, concise, and exact terms so as to enableany person skilled in the art to which it pertains, or with which it ismost nearly connected, to make and use the same.

What is claimed is:
 1. A metal colloid monolayer comprising a pluralityof colloidal Ag clad Au nanoparticles surface confined on a substrate,said monolayer having a gradient of nanoparticle density in a firstdirection, and a gradient in particle size in a second directionperpendicular to said first direction.
 2. The metal colloid monolayer ofclaim 1, wherein said nanoparticles are bound to said substrate bybonding with functional groups of a bifunctional organic film.
 3. Themetal colloid monolayer of claim 2, wherein said bifunctional organicfilm is selected from the group consisting of an organosilane,poly(allylamine) hydrochloride and biotin.
 4. The metal colloidmonolayer of claim 1, wherein said substrate is elected from the groupconsisting of glass, quartz, alumina, tin oxides and metals.
 5. Themetal colloid monolayer of claim 1, wherein said nanoparticles have asize of about 3 nm to about 100 nm.
 6. A metal colloid monolayercomprising a plurality of colloidal Ag clad Au nanoparticles surfaceconfined on a substrate, said monolayer having a gradient ofnanoparticle density in a first direction, and a gradient in particlesize in a second direction perpendicular to said first direction formedby: coating a substrate with a bifunctional organic film; immersing thecoated substrate into a colloidal Au solution such that a first leadingedge of said substrate is immersed in said colloidal Au solution for alonger period of time relative to a first trailing edge to provide amonolayer of Au nanoparticles having a decreasing level of coverage fromsaid first leading edge to said first trailing edge; withdrawing saidsubstrate from said colloidal Au solution; rotating said substratethrough an angle of about 90°; immersing the substrate provided with themonolayer of Au nanoparticles in an Ag⁺ solution such that a secondleading edge of said substrate is immersed in said Ag⁺ solution for alonger period of time relative to a second trailing edge to provide amonolayer of Au nanoparticles having a decreasing level of Ag claddingthickness from said second leading edge to said second trailing edge;and withdrawing said substrate from said Ag⁺ solution.
 7. The metalcolloid monolayer of claim 6, wherein said nanoparticles are bound tosaid substrate by bonding with functional groups of a bifunctionalorganic film.
 8. The metal colloid monolayer of claim 7, wherein saidbifunctional organic film is selected from the group consisting of anorganosilane, poly(allylamine) hydrochloride and biotin.
 9. The metalcolloid monolayer of claim 6, wherein said substrate is selected fromthe group consisting of glass, quartz, alumina, tin oxides and metals.10. The metal colloid monolayer of claim 6, wherein said Aunanoparticles, prior to cladding, have a uniform size of about 3nm toabout 100 nm.
 11. A method of determining optimal surfacecharacteristics of a metal colloid monolayer comprising a plurality ofcolloidal Ag clad Au nanoparticles surface confined on a substrate foruse in an analytical procedure, said method comprising: coating asubstrate with a bifunctional organic film; immersing the coatedsubstrate into a colloidal Au solution such that a first leading edge ofsaid substrate is immersed in said colloidal Au solution for a longerperiod of time relative to a first trailing edge to provide a monolayerof Au nanoparticles having a decreasing level of coverage from saidfirst leading edge to said first trailing edge; withdrawing saidsubstrate from said colloidal Au solution; rotating said substratethrough an angle of about 90°; immersing the substrate provided with themonolayer of Au nanoparticles in an Ag⁺ solution such that a secondleading edge of said substrate is immersed in said Ag⁺ solution for alonger period of time relative to a second trailing edge to provide amonolayer of Au nanoparticles having a decreasing level of Ag claddingthickness from said second leading edge to said second trailing edge;withdrawing said substrate from said Ag⁺ to form a monolayer having agradient of nanoparticle density in one direction, and a gradient inparticle size in another direction; using the gradated monolayer for ananalytical procedure; and analyzing the results achieved at differentregions of said gradated monolayer to determine a nanoparticle coverageand particle size that provides an optimal result.
 12. The method ofclaim 11, wherein said monolayer is a surface enhanced Raman scatteringresponse substrate.
 13. The method of claim 11, wherein said monolayeris a biosensor for detecting the presence of a biological ligand.